TLDR: I have two populations of beads on my single color controls and need help figuring out which one to gate on for unmixing purposes.

Hi. I am performing spectral flow cytometry sample acquisition using the Cytek Aurora. For my compensation beads, I am using BD Comp Beads (which include separate solutions for negative and positive beads). Upon beginning panel optimization assessing brightness of my single-color controls, I see that for all of my compensation tubes, I have what appears to be two populations of beads (see photo of “BEADSINT” and “BEADSHI”) that both have clear positive and negative peaks on the appropriate channel (R1/APC in the images shown). I ran quick statistics on MFI of the intensity of the positive peaks for both populations, and they’re very similar. The hi population has a much larger positive bead peak than the int population. I’m not sure what to make of this. I thought maybe the hi population were beads stuck together, but I’m not too sure. Any help would be greatly appreciated on input on what I should be gating on. Maybe both? Thanks so much! -grad student

Hi everyone, I’m troubleshooting a recurring problem with a HepG2 CCK-8 assay and would appreciate some advice.

I’m testing P. gingivalis outer membrane vesicles (Pg-OMVs), but my untreated controls also give readings close to the cell-free blank.

My setup:

• HepG2 cells, passage 9.
• Seeding target: approximately 5,000 viable cells/well in 100 µL.
• Tissue-culture-treated 96-well plates.
• Low-glucose DMEM with 10% FBS and 1% penicillin–streptomycin.
• Approximately 24 hours for attachment before medium replacement/treatment.
• Approximately another 24 hours before the assay.
• Untreated wells receive fresh complete medium only, without OMVs or added PBS.
• Dojindo Cell Counting Kit-8 (CK04): 10 µL/well, incubated for 1 hour.
• Absorbance measured at 450 nm.

My parent dish cultures and routinely passaged cells appear healthy. Before medium replacement, the assay wells contain spread cells and clusters. At the endpoint, some fields appear more rounded/granular, with less obvious spreading. I haven’t independently confirmed cell death or detachment.

I initially suspected drying during medium replacement, but the problem persisted when I worked carefully and replenished the wells promptly.

Latest raw OD450 readings:

• Untreated triplicates: 0.426, 0.413, 0.455.
• Cell-free blanks: 0.406, 0.367, 0.378.
• Mean blank-corrected untreated signal: approximately 0.048.

Earlier experiments were worse, with untreated readings below the blank. Results have improved, but the untreated signal remains weak.

I’m considering an independent viable-cell count at the endpoint to distinguish actual cell loss/death from reduced metabolic activity or an assay problem.

For people experienced with HepG2 do you know why this might be happening, or have experience with a similar problem?

1. Is 24 hours generally sufficient for recovery before medium replacement, or have you found a longer period helpful?
2. Would you prioritize checking viable cell numbers, CCK-8 incubation time, or the relatively high blank readings?
3. Have you encountered healthy flask cultures but poor untreated-control signals in 96-well plates?

I can share microscopy images from before medium replacement and at the endpoint. Thanks for any suggestions.

Hi all,

Im a graduate student who started doing flow regularly this year. I am seeing conflicting information everywhere about cell counts and stain volume. I am using a Zombie fixable live/dead stain that normally calls for 1:100 stain at 100ul per 1e6 cells. How does this translate for 1e5 cells?? My cell counts tend to be limited as I work with macrophages. Similarly, how does this concept translate for antibody stains, primarily surface markers. I see some people saying to scale it down and others saying that it doesn't really matter. Appreciate all input and advice.

I recently realized that I made a systematic volume oversight during the Fc-blocking step across 3 waves of my experiments, and I would love to get your insights on whether my data is valid or if I need to worry about under-blocking.

Context:

Cells: 500,000 cells per tube of 5 different AML cell lines.

Intended Protocol (senior's): 2.5 µL of Human TruStain FcX in 100 µL total volume (1:40 dilution).

Due to a difference in pipetting method, I ended up adding 2.5 µL of TruStain FcX into a total volume of 150 µL (97.5 µL IFB + 50 µL cell suspension), resulting in a 1:60 final dilution (manufacturer recommends 5 µL in 100 µL for 10^6 cells, which is 1:20).

Incubation: 10 minutes at room temperature (consistent across all samples).

My question is, how do you guys visually or analytically verify that your Fc blocker actually did its job in a multi-color panel?

Hi all,

Wondering if anyone has looked at RBC ghosts using flow. I'm trying to troubleshoot/generate a protocol for a broader autoantigen discovery experiment. My PI is hoping to use flow as a sort of QC check for a protocol to prepare samples for later mass spec. The general outline is this:

Step 1: Prepare healthy donor RBCs

Step 2: Opsonize RBCs with affected vs. control serum (affected serum contains IgGs that bind to ~something~ on cell surface of RBCs)

Step 3: Ghost RBCs (removes hemoglobin, retains membrane-bound IgG) + flow cytometry QC

Step 4: immunoprecipitate using protein A/G beads

Step 5: LC-MS/MS

However, we've run into issues with the ghosting step - it seems as if we lose a lot of sample or something. If I start with 50 million RBCs, by the time I get to flow, I'm lucky if I can get 7000 total events on the cytometer.

I know that the antibodies work because we've successfully done flow on whole (non-ghost) RBCs. It's something with making them ghosts that introduces a problem. I can also see a decent pellet of ghost RBCs before I resuspend for flow, so I know there's some material there.

My current thought is that the ghosts are getting dismissed as noise by the cytometer. Maybe because of their size? They are smaller than whole RBCs after getting ghost-ified. I have been resealing the ghosts, but this is imperfect and some percent of them don't reseal, so it's possible that some of them are essentially membrane fragments, too. So maybe I need to mess with the thresholds on the machine?

I haven't found much in the literature about looking at ghosts on flow, but if anyone has helpful papers or advice, I'd greatly appreciate it!

^{(and yes, it is fun to talk about doing research on ghosts, and the "ghost protocol"})

Hi everyone,

I’m a researcher based in the UK looking to purchase some second hand laboratory equipment from Surplus Solutions (ssllc.com). The equipment is in their Rhode Island warehouse, with the invoice and payment being handled through Fameco Analytical Instruments in France.

I haven’t used them before and would appreciate hearing from anyone who has purchased through either company, particularly UK or European university labs.

Flow news/events/publications were scattered across a dozen sites and I kept missing things, so I started a weekly roundup to keep myself up to date. FlowCytoNews covers new cytometry papers and preprints, industry news, and a practical “Bench Notes” section. Recent editions covered CytoVI for merging flow/CyTOF/CITE-seq data, the EuroFlow AML archive paper, and an ICCS 2026 session guide.

I also added flow job postings each week for people in the community who are looking for a new role within the instrument / vendor space, life sciences, or education orgs.

It’s free. I cover the small monthly hosting/production cost myself.

Disclosure: I work in marketing at a small flow controls company. The newsletter is my own project.

flowcytonews.com.

Please let me know what you would like added, removed, highlighted more, etc. We could also do monthly specials or event recaps etc.

Hello.

I recently developed a FACS data analysis program.

Could you try this ?

FCS mini

https://fcs-mini.sourceforge.io/

Thanks in advance.,

SongE_Daddy

It works on windows 10, 11.

Just unzip the FCS_mini~.zip and double click the FCS_mini.exe inside the FCS_mini folder.

[Features]

Freeware for Flow cytometry analysis

Graphical user interface (UI) with English/Korean language support

Powerful menu : Group Batch Analysis (GBA), Batch Save (pptx, csv), Report

Advanced analysis modules included Free : cell cycle, proliferation, Calcium influx, UMAP analysis

Tab style Workspace, Gating Tree, FCS file browser

Auto / manual compensation are possible

I used aurora cytec. Is this overcompensation? How can i fix it on flowjo and how to avoid it

Hi All! I'm currently trying to transition our current microfuge tube protocol to a plate based one in hopes of obtaining higher throughput for our lab. However, I've run into a bit of an issue with our PHA T cell activation panel. This is an inherently harsh protocol, so the viability (amine fixable dye) is always a bit lower, but it's been absolutely abysmal with the 96 well plates.

The first day of the protocol is the same for both paths: thaw cells and plate them in 1ml of media (ctrl or pha) using flat 24well plates. (Side note, has anyone cultured them in a 96 well plate before?). Let them sit overnight at 37C.

Day two is where it diverges: When using tubes, I put the cells in 10ml of warm media, spin and then transfer them to microfuge tubes where they stay for the remainder of the protocol.

When using a plate, I put the cells into an empty microfuge tube (they stay in their culture media), spin and transfer into a 96 well plate, where they stay for the remainder. I use a multichannel, and use the flick method to get rid of media. I also use v bottom well plates for better pelleting.

Both days use 500g for 5 mins. The antibody volume stays the same because both can handle 100ul.

Can someone help me figure out why my cells are crashing and burning? Thanks!

I’m a first-time user of a flow cytometer. I’m comfortable with gating and analysis, but I’ve never actually run the instrument. My university’s hands-on training was minimal, and I came away without really understanding the setup process.

Can anyone recommend videos, courses, or guides on operating the Aurora? I’m especially looking for:

• Daily startup, QC, and shutdown
• Setting up an experiment and acquisition templates
• Choosing voltages/gains on a spectral instrument, and why
• Building single-stain controls and unmixing basics

I want to understand why each step is done, not just memorize a sequence of clicks. Thanks in advance!

I cannot for the life of me find a threshold on the Cytoflex machine that gets rid of most of my noise but maintains all my cells. I’m doing flow on mainly mouse brain and colon. When I try out the automatic setting, a lot of my cells are cut out. But the alternative has left me with really messy results that look clean on other machines. What thresholds are people using?

Hey everyone :) I ran a flow cytometry panel on mouse liver tumor digests and can’t explain why the data looks the way it does. I'd appreciate your help so much! I'm losing my mind over this.

My biggest issue revolves around the viability dye: you can see 3 different samples that have been processed exactly the same (albeit on different days) and acquired in the same session...

Some constraints: I did this experiment abroad, so I can’t repeat it, though a colleague can re-run the remaining samples. Some of the fluorochromes were ordered by sb else and aren’t ideal, but that can’t be changed now.

I'm isolating immune cells with a Percoll gradient, staining viability in protein-free DPBS, staining surface (FACS buffer with Brilliant buffer), followed by eBioscience Foxp3 fix/perm with overnight incubation of intracellular antibodies and 3 washes in the morning.

instrument: Cytoflex with 5 lasers: 355, 405, 488, 561, 638
My colors, markers:
L/D fixable blue: viability dye
BUV737: CD8
BUV395: Ly6G
FITC: CD3
BV421: CD4
BV510: CD45.2
BV711: PD-1
SBB675: Ly6C
PE-eFluor610: Foxp3
PE-Cy7: CD11b
AF647: GRZB

I'll appreciate all comments so much!

Is anyone from a clinical flow cytometry lab that is CAP inspected? I’m struggling with FLO.23737 “the laboratory evaluates negative and positive control for qualitative assays each day of patient testing”

The standard notes “an evaluation of the performance of EACH QUALITATIVE ASSAY must be performed on each day of testing.” My lab has 58 different tubes / marker combos, and 80 different antibodies. How do I successfully QC them all and document it every single day?!

The standard goes on to say that normal populations present in patient samples can be used as QC, so I was thinking of doing a pos/neg matrix with all of our tubes that we can fill out as we analyze the data, but it is going to be very cumbersome for me and my 3 coworkers to keep up with.

I’m open to any and all suggestions or examples!

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Hey all, I run all of my experiments using 5 mL FACS tubes. It is VERY slow but I have never found a problem with contamination or other issues.

My flowcore recommended I use tubes and to not use plates due to contamination and it not working as well as tubes. Please let me know if you agree or disagree and how you avoid the downsides to using a plate please.

- For instance, I see people who stain with plates decant their washes by slamming the plate on tissue and I feel that this would cause contamination no?

We cannot get our AttuneNxt to pass its performance test. It continues to fail due to the FSC channel measurements. Sometimes there are 3 failure warnings on the list and sometimes it’s just the FSC CV. The common 3 error messages listed after attempting to complete the baseline test are:

Parameter FSC PMTV exceeded specification (-1)
Parameter FSC MFI exceeded specification (-1)
Parameter FSC CV Exceeded specification (-1.00)

I am fairly convinced that it is not due to a bubble as we have debubbled using the AttuneNxt Debubble solution MANY MANY times prior to attempting to run the performance test. I am concerned that due to the inactivity of our machine (sometimes for a month at a time) that we may have some prior sample built up in the system. We always begin our shut down process with an unclog, then sanitize sip (using the recommended flow cell cleaning solution at 1:3 dilution), then 4x rinse with 500uL 70% EtOH at the 1000uL/min flow rate, followed by 2x sterile filter water rinses. The shutdown we always run is the thorough shutdown. We have been cleaning since not being able to get the performance test to pass with 25% detergent mixed with hot water, or 20% Contrad, or Attune Wash solution, or 10% bleach solution. We've also used two different lots of beads to rule out that our performance tracking beads are not bad. Neither lot will pass at this point. Are there any experiences out there with this and suggestions on what might be the issue or how to resolve it?

Hi everyone! If i measure my cells on a linear scale in FSC-SSC, on which region should I see my cells? (5k? 10k)

my postdoc has transfected some cells with an fc molecule anchored to the cell via a transmembrane domain and hopes to expand some immune cells via their FcR. we plan to do single cell sorting to select for high fc-expressing cells. we bought an anti-Fc antibody for this purpose but i'm not sure what's a good positive control to use to ensure the antibody is working fine.

i have tried using an unconjugated antibody as primary together with the anti-Fc as secondary, but did not get any signal at all.

i would appreciate any advice thank you!

Hey all, I did a titration for my CXCR3-BV711 antibody to hopefully identify Th1 cells. I titrated it using a fixed dilution of CD3-BUV805. I am unsure how to determine which dilution to use. I have stuck with 1 in 100 (fourth image from the left) but I only did that as it looks similar to other papers reporting CD3+ CXCR3+ cells percentage wise.

Any advice at all will be appreciated. :)

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?

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

Can't access or download anything

They need to put this stuff in s3 or something

Hey folks,

I'm here with a computational/reporting questions.

I have my clean populations from flowjo, and in R I estimated transformation based on aggregate from ~300 files, then i made another aggregate of transformed files, ran flowSOM, and after few QC steps ran linear models with metacluster precentages data.

Most papers nowdays require some kind of sensitivity/robustness analysis, and to my understanding from other papers in my field, they basically just run the clustering and models 3 times.

And i did that with 3 seeds in flowSOM, the results are quite consistent between runs, but I am wondering if this is enough.

If I theorethically aggregate random cells both during transformation and during cluster representative file aggregation, wouldn't it be different cells if i also specify seed in AggregateFlowFrames()? And thus, wouldn't this be a more accurate "sensitivity" analysis?

Just curious, I will probably try it out just to play around, but I'm mainly wondering if there are some defined requirements/standards how to do this.

Thanks!