Hi Melissa,

I have experienced problems with cell adhesion in 96 well plates too, although not whilst developing an ELISA. My cells were Caco-2 monolayers, and liked to adhere closely with each other, as well as sticking to the growth surface. However, during wash steps with PBS, I found the edges of the cell clusters would start to peel off the growth surface, and due to the close adhesion to the other cells, this would cause a high loss of cells from my assay. This only occurred in 96-well plates, where the growth surface area was small. I don't know why PBS caused this, but I got round the issue by washing in serum-free growth media and manual aspiration with a pipette, both seemed to help. It would certainly be worth a try?

Best of luck,

Mark

Hello Melissa,

Your cells are dying obviously... Do you use pre-coated plates? This might solve your problem.

Karin

Hi Melissa

1% Hydrogen Peroxide does seem a very high concentration. I would suggest removing the step and see if the cells are happy. Then I would start adding lower conc of peroxide if it is essential. Maybe start with 0.1%. Fixing with ice cold methanol may also be an option. Good luck.

What type of cells do you use?

Another way to remove the liquid from the wells could be using a pippete or a faucet suctor. Be sure to incline the plate a bit and avoid contacting the cell monolayer with the tool yuo selected.

What cells are you working with?

Increased autofluorescence after paraformaldehyde fixation is often the result of incorrectly made solution. You could try glutaraldehyde instead. I would also agree with Shailendra that without fixation 1% hydrogen peroxide will probably be killing the cells, which sound as though they are dying rather than detaching.

Hope this helps.

Hi Melissa

I faced the same problems with immunostaining but I over come the problem by:

I think in this case it is better not to leave the cells reach 80% confluence because the will make the cells easy to peel off during the staining it is very important from experience to grow the cells in low number like 50%

The second important think is to wash the cells manually and carefully

Avoid harsh technique try using simple washing by multichanel micro pipette

Cell number is very important so keep it low to avoid the cells from come out

Best Wishes

F

I'm using Osteoarthritis fibroblast like synoviocytes

I agree with Keith: the hydrogen peroxide is killing your cells. Fixing them (as you did) will kill them too, but leave them attached to the plate in approximately the shape and size they were when alive, so you should think about modifying your protocol to include fixation and then cope with the background problem. Bone cells are notoriously detachable, and I feel fixation should be included in your protocol.

The step of 'adding peroxide to limit engogenous HRP activity' is absolutely necessary, but only if your antibody system uses HRP enzyme activity to work. You could switch to one that has fluorescence as a visualization marker ... but that will come with its own "background" problems.

If you switch to including a fixation step, you will need to recalibrate most of the parameters of your antibody usage (concentration of primary, concentration of secondary, timing, composition of blocking buffers) to optimize signal-to-noise, and you may want to include new steps, such as a rinse step with a background dimming buffer applicable to your visualization marker.

Pierce has a page on how to reduce background that might be helpful: http://www.piercenet.com/browse.cfm?fldID=5E97EE7E-99FE-A688-C8BC-D15A791DBA4A

Hi Melissa,

I agree with all the suggestions and comments regarding probably toxicity with 1% H2O2 and the potential use of fixatives to help with cell adhesion.

I would exercise caution however when using fixatives such as 4% Formaldehyde, glutaraldehyde or methanol, because these agents can lead to some permeabilization of your cells. And as I understand the question you are asking regarding ligand binding (to the cell surface only?), this could be a potential confound in the results obtained.

You can refer to this discussion where several comments suggest that fixatives are sufficient to permeabilize cell membranes: https://www.researchgate.net/post/Does_formaldehyde_permeabilize_the_nuclear_membrane

Link for article on immunolabeling artifacts for all interested:

http://www.nature.com/nmeth/journal/v9/n2/abs/nmeth.1855.html

Best of luck

I agree with all points mentioned above: about the H2O2 step: I would suggest removing it altogether and just be sure to keep the control (untreated cells) to see what endogenous HRP does in this system. Regarding the coating, maybe use some collagen or anything else (poly-D-lysine?) to coat the plates and be as careful as possible with manual aspiration and inclination of the plate, after centrifuging the plate with each step. Maybe as a parallel approach, also wash the plate as you would normally, collect all washes separately in another plate, spin those separately and add back those cells to the respective wells in the first plate. That way, you won't lose loosely adherent cells.

I do this type of experiment using HEK cells to detect surface receptors. I have to plate cells in dishes pre-coated with poly -D- Lysine. Some use collagen coated plates. If I do not use these coated plates, my cells will detach just like yours do.

John

Hi Melissa -

I have run cell-based ELISA using a variety of cell types, some adherent, others suspension. To keep the cells attached to the plates, I coat the ELISA plates with 100 ul/well polylysine solution (10 ug/ml) for 1 hour at 37 degrees C, wash with PBS, then add the cells at 100 ul/well (5x10^4 cells/well). After allowing the cells to settle for about 4 hours at 4 degrees C, add 100 ul/well of a 0.5% glutaraldehyde solution (made in PBS). This will cross-link the cells to the plates. Wash plates gently (no plate washer) and proceed with the ELISA.

Good luck,

Ed.

Poly-L-lysine may help to stick down cells, as noted by others. Note that it can also change the behaviour of your cells, something you have to control for at first. Hydrogen peroxide is certainly toxic and should be eliminated. Gentle washing can be achieved by repeatedly immersing the plate by hand into a container of buffer to flood it and then flicking out the buffer into a sink to remove it. Even a hand-held wash bottle is gentler than an automated plate washer. Blot and wipe afterwards to remove excess liquid before proceeding to the next step. Centrifugation is unnecessary, certainly during washing! In any case you don't need to use 1000g, which is probably also harmful to your cells. 100g is more than enough, but why do it at all? If your reagents are not flowing evenly into the wells it is better to shake the plate gently than to centrifuge. You also save valuable time. Does your PBS contain Mg and Ca? If not the cells will certainly detach. Also, it is a good idea to add glucose to keep the cells happier. Everything should be cold too, or the cells will pino/endocytose or cap your immunoreagents. Time is the most important factor because the cells are under serious stress during the whole procedure. Optimize all incubations to reduce the time to the absolute minimum. You'd be surprised how fast those reactions really are.

I tried eliminating H2O2, but the cells are still dislodging at a later stage in the experiment. I have been using PBS without calcium and magnesium to wash and to mnake 1% BSA to block and dilute my peptides in. I have tried fibronectin.

If you add 300 ul of wash buffer, do you aspirate all the 300 ul out?

Hi Melissa. One thing that could be contributing to the problem is the use of calcium/magnesium-free PBS. Cells need these to stick down and can very quickly lose adherence without these ions (that's of course why we specifically use this PBS for trypsinisation! Try using with calcium and magnesium or switch to HBSS.

To repeat, your most serious mistakes are to use hydrogen peroxide on live cells and to use calcium/magnesium-free PBS for washing and blocking. Additionally you are wasting time centrifuging (and centrifuging at excessive force), and time is everything in this type of assay. There is no reason to aspirate, which can also dislodge cells. Do the flood/flick method of washing, which all IHC/ICC students eventually learn (reluctantly, it seems). We use it frequently in ELISAs, even to the point of taking plates offline from a robotics platform and doing the flood/flick washes manually, then putting the plates back online. In some cases this improves variability by an order of magnitude! Don't knock the low-tech approach. If you don't know what I mean, here it is in detail: (if you are right handed) Hold a wash bottle of cold buffer in your left hand (cut off the tip of the dispensing tube so that the dispensed stream is broad and therefore gentler). Hold the plate in your right hand over a sink. Flood the plate with buffer from the wash bottle. Don't worry about volume, the plate and all the wells in it should be overflowing. Turn the plate over and flick the contents forcibly with a flick of the wrist into the sink. Repeat at least four times. For the last wash flick several times into the sink to remover all liquid then blot the plate on a layer of tissues to remove all adherent liquid. You can do half a dozen washes in a minute. When you do the statistics, it beats the hell out of automated washers, not to mention that your cells will still be there. Believe it!

I've done a lot of IHC and use the flood/flick method for washes. It works! Try it.

Hi we are trying to perform a cell based ELISA and are having the problem of very high background and cell detachment. We fix our cells with 4% PFA for 30 mins at 4 degrees. Since we are trying to detect expression of a cell surface protein we omitted the quenching step, as in our experience it makes the cells excessively permeable.

Could someone kindly share a CBE method for detecting cell surface protein which would help us get around this isuue