Hi Filip,

I assume that you're using a commercial antibody, do they specify on the data sheet the best fixation for this antibody? Otherwise you can try paraformaldehyde 4% in PBS, 20 min at room temperature, which is standard. Do you add the triton in the fixative? This might cause the leaking from the cytoplasm and the cells to look unhappy. After fixation you can try to permeabilize with 0.04% saponin which is milder than triton and should cause less cytoplasm leak (what is your triton %?). Permeabilization with saponin is reversible so you will need to keep it in the blocking buffer and with the antibodies.

Also the autofluorescence might come from the glutaraldehyde (did you observe it with ethanol fixation?). You can quench it with 0.1% NaBH4 (sodium borohydride) 3x1min, after fixation. The autofluorescence with PFA should be lower.

You can also fix with methanol, 20%, 6min at -20C. That will also permeabilize your cells.

Hope it helps!

oups! sorry, not 20% methanol!!!!! but pure anhydrous methanol cooled down at -20C prior to fixation!

Anti-Myc-tag mAb-Alexa Fluor® 488's protocol advises to fix the cells by immersing the slide in PBS containing 4% paraformaldehyde (PFA) for 15 minutes at room temperature. That's what we did - except we didn't have a fresh paraformaldehyde so we used another cross-linking aldehyde - glutaraldehyde. Following that protocol all samples were destroyed with only a few cells in good shape. The autofluorescence was observed in both ethanol and aldehyde method. It can't be the antibody, because we always include a control sample stained with everything except for the antibody. It was about 2x stronger in H9C2 than in CHO cells.

Is there a difference between methanol and ethanol fixing? Ethanol fixation we did: 100% ethanol for 15 minutes at -20C.

There is also one more drawback - from what we've seen ethanol fixation decreased cells' size by about 50%.

Usually ethanol is used in combination with glacial acetic acid or with methanol. I tried ethanol vs methanol alone and I had better results with methanol. However if the protocol says 4% PFA I think that this is what you should try. Some antibodies might work really well in a fixative and not at all in an other.

Alcool fixations dehydrate the cells which can explain that you cell's size decrease. For example with plated skeletal muscle fibers methanol fixation is terrible, they shrink and get distorted.

An other possibility is also that your transfection did not work. Do you have an other way to test it? By western blot or by using a fluorescent construct?

Hi Filip,

Best advice is don't change method. Formaldehyde and glutaraldehyde while sharing similar structures are really best suited to different applications. Glutaraldehyde will give more auto fluorescence and more cross linking which might impair antibody staining. The ice cold methanol advised above is a good option to try if 2-4% formaldehyde doesn't give suitable results.

Thanks for your answers

Hi Filip, are you sure that your H9c2 cells are really transfected. I tried to transfect this cell line and it is not easy. Lipofectamin leads only to transfection efficency of

Hi Filip. I've used

Hi

Andreas - I will be testing the transfection efficiency again, with Western blot and Real-time PCR. I will use both CHO cells, which are said to be transfected very easily, and H9C2 cells. We have no idea if the antibody staining works - after we confirm the expression we will see if the method for fixing and permeabilizing works well with the antibody. This is the official procedure recommended by the manufacturer:

http://www.cellsignal.com/support/protocols/IF.html

Last time we followed it, we could see about 1% transfection efficiency and a strong signal from our antibody.

As for H9C2 transfection - we use Lipofectamine 2000 and gave Fugene HD a try, but without much success. Did you manage to solve this problem?

Nicholas - yes, we are using exactly those chamber slides coverglass, the glass itself doesn't give any fluorescence. It's only within the cells. As I recall, the blocking solution contained BSA, but the antibody is mouse. I wonder if blocking with mouse serum would reduce the unwanted fluorescence as I think the antibodies can be bound nonspecifically in our cells. I'll look if the fluorescence looks like signal from unwanted antibodies (Alexa Fluor 488 emission).

Another consideration is to change your blocking solution. With a particular antibody I used before, I blocked with 1% BSA but it did not work, but when I used 5% normal goat serum (1h, room temperature) my antibody worked well. You may need to optimised this (some people do up to 10% NGS). And some people do a combination of BSA and NGS.

Also my experience with the chamber well slides is that 100% methanol fixation causes the chamber wells to dry and become frosted/opaque. Therefore the antibody solution may flow between the wells, which means that your negative control (that you thought had no primary antibodies might in fact have received primary antibodies from the well next to it).

I hope some of these considerations help. Also it will be best to stick to PFA if that is what has been recommended by the manufacturer.

I also recommend using a nuclear stain like DAPI or Hoechst to ensure that you know where the signal is localized.

Also more importantly what is your secondary antibody raised in? I don't think that there are too many issues regarding your use of a primary antibody raised in mouse being used on mouse cells (happy to be corrected). I normally have secondary antibodies raised in goat (so I would have a Alexafluor 488 goat anti-mouse secondary antibody).

The antibody is from mouse, the H9C2 cells are from rat. We blocked with BSA because we didn't have any mouse serum around. Haven't tried goat serum. The antibody is a primary antibody, with Alexa attached to it so there are no secondary antibody issues (fortunately!). Additionally stained with DAPI and with red ATTO phalloidin. Those look excellent.

Oh, so is it not necessary to block the tissue with the same species serum as the antibody? BSA or NGS can block well when using mouse antibodies?

This website may be useful which has this line "A critical factor, though, is to use serum from the species that the secondary antibody was generated in, as opposed to the species of the primary antibody." http://www.piercenet.com/method/blocking-strategies-ihc

I haven't used a conjugated primary antibody before, so I'm not sure. Maybe you do need mouse serum? My guess is to use mouse serum (but hopefully someone else might answer your question).

Hi Filip,

I use jetPEI, Lipofectamin 2000, jetPRIME, Fugene and have the same problems. Also electoporation is no very good. For stainings it is ok, but for Western Blots it is difficult. It depends then deffnitely on the protein leven of the transfected cells. If you find a high transfection method which really works in your hands, let me know. I wish you mutch success. Best Andreas

Hi Filip,

I use 2 % paraformaldehyde (15 min) and after washing, 0.1% triton x-100 (20 min).