I am trying to determine antibody concentration of immunized mice by incubating transfected HEK cells with mice sera. As I have many samples, it should be established in a 96 well plate. I detach the cells by incubating them with EDTA and resuspend with PBS. Then, cells are centrifuged (1000 rpm, 5 min) and the pellet is resuspended in FACS-Buffer (1% FCS, 0,1% NaN3 in 1xPBS). Afterwards, I transfer 20 µL into 80 µL FC-Block-solution in FACS-Buffer (Probably, I can omit the FC-Block for HEK cells) in each well of a 96-well-U-bottom cell culture plate (polystyrene). Then I add, the sera, incubate, centrifuge at 1700 rpm, 5 min and discard the supernatant by performing a gentle but firm swing with plate up-side-down into a sink. Then I resuspend with 200 µL PBS let it sit for 5min and centrifuge, discard, incubate with 2nd AB, centrifuge, PBS, discard and resuspend in 200 µL FACS-Buffer. At the end, there are only a few cells left. And as the transfection rate is also quite low (
We use V-bottom plates instead of U-bottom (NUNC, come in packs of 4 with no lid, lid ordered separately), and we always spin plates for 10 min, and I found 5 min to cause some cell loss as well.
I bet FcBlock is just an expensive non-specific protein block in your case :)
Yes, I always use V bottom plates; generally leave an empty well between samples; wash 1 or 2 times only with 200ul of PBS; centrifuge at 1200rpm for 2min (some cells will die at higher speed); do a single swing over the sink (about 90% of the PBS is gone leaving about 90% of the cells).
Good luck
Thank you Eric for your answer! I am wondering if a swing is better than removing the supernatant by the use of a pipette. I have the feeling, I have more control then..
Thanky you John! To my knowledge, sodium azide is not mainly used as a preservative but rather as a metabolic inhibitor that presents internalization of surface antigenes. But it is a good point you mentioned with cell viability. My cell viability is
We use polypropylene V-bottom plates in our lab. It reduces the risk of adherent cells to stick to the plastic surface.
Okay... My cells are dead at the end of the staining.. My protocol: 100 µL cellsuspension of a 5,8*10^6/mL Suspension in FACS-Buffer (fresh, with 10%FCS and 0.1% NaN3) per eppendorf tube. 1h incubation with sera. centrifugation (300g 10 min 4°C). remove supernatant with swing and drop it on a tissue. add 200 µLFACS-buffer and flick the tube over a grid to dissolve pellet. sit for 5 min. centrifugation. discard supernatant add PBS, flick, let it sit for 5 min. Centrifugation, discard, 2nd AB in FACS buffer, flick, 30 min. centrifugation, washing step as before.
I compared a control (without serum) with the original population I used for the staining. And the vital population is just lost.. Too many washing steps? Flicking bad? Should I add EDTA?
Hi Betty,
as mentioned above there are several possible pitfalls here. For Cell loss: If you are testing right now in Ep-tubes with many cells you will always see the pellet afetr fuging and will be able to see cell loss/pellet distortion. So you can go an try 3mins at 300g, which should be enough for a stable pellet. you can do side by side swing an pipet off method. To increase starting cell number for the experiment you could try 2 48-well plates, and transfer to 96-well after trypsinisation.
Viability: Additionally I would go and start of with 10 (orso) identical tubes and check viability after each (major) step in the prot. Like after each fuging step and maybe after resuspendion/before next fuge.
Try one tube without NaN3.
Viel Erfolg!
Hey everybody,
here's an update: Within our original population in the flask, we have about 30% positive HEK cells (P3).
My supervisor repeated the experiment and used a FACS-buffer without NaN3. He had the idea that our filtration step at the end prior to measuring is the problem. Within our first experiment, we used mesh from FALCON (40 µm) and had a nice population. But even if we used 100 µm mesh, we didn't see a population this time.
Within our first experiment, I also forgot to incubate cells with sodium butyrate (usually we incubate cells with 10 mM Na-butyrate for 16-20 h before starting the staining). So we didn't incubated them with Na-Butyrate this time. Then my supervisor washed cells of with PBS (1), trypsinized the rest, stopped reaction with PBS (2) and splitted this solution (2a). The latter, we fixed with ice-cold MetOH, so that we had 3 different populations to check.
Under the microscope, 1 had almost no positive cells, 2 had about 5-10% and the 3 about 5%. All of them were quite weak in the signal though. We went directly to the FACS but there was no poulation in all of the samples.
I re-checked the sample under the microscope: in all the three sample we used for FACS were no positive cells dectable. In the original samples (without filtration), I could see at least some weak positive cells in the MetOH-fixed samples. Then I re-checked a PFA-fixed sample from the 19th of January and I could clearly detect positive cells in there which were also quite strong in signal.
This sample was the original population, unstained, unfiltered, which had unterwent Na-Butyrate incubation, trypsinization, and have been kept in FCS-Buffer with 1% FCS and 1% NaN3. To me, it seems that the filtration with our mesh is a problem but incubation of FACS-buffer with NaN3 seems to stabilze the cells or at least our protein. Maybe, they need more stabilaztion (Buffer with 10% FCS).
As I still have cells, I want to do retry it with 5 different buffers:
1) 10%FCS, 1%NaN3
2)10%FCS, 0,1% NaN3
3) 1%FCs, 1% NaN3
4) 1% FCS
5) 1% FCS, 1% NaN3, PFA fixed.
What do you think?
I want do trypsinize cells and stopp reaction with PBS with 1% FCS
Hi Betty,
I think you can get better result if you use 96-well-V-bottom (but not U shape) plates. I use 1700 rpm and 2 min and on my hand it is very good to overcome cell loss.
Indeed when you use swing out rotor, the tubes turn 180 degrees and this helps the cells to form a nice pellet instead of smearing down the tube wall.
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