Hi Priya

Without sharing your protocol or images of the non-specific staining it's a little tricky to give any substantial feedback sorry. In saying that, my bet would be that the background is almost certainly not to do with your cells, but most likely something simple with your staining protocol. I've performed IF on 293s many times and never seen any background staining in my controls. There are several sources of high background during a staining protocol, these include: using the wrong block solution, using too much antibody (especially secondary antibody), allowing the coverslips to dry out during the staining process, not enough washing (or not stringent enough, ie no detergents), and contaminated (anti-fade) mountant. There are many other reasons for high background, but these are probably the most common.

Hope this helps.

Good luck

Software to acquire microscopic images tries to show signals, no matter how the signals are low when you use an automatic setting. Take pictures of cells you think are expressing your interested proteins first and save or take a note of the condition you use. And try to use the same condition to take pictures of the negative control. If you are not aware of this, check the manual of the software you are using.

If you did not use detergent in your buffers for your staining, you might want to add 0.1%TritonX100 or Tween 20/80.

By the way, you do not see the endogenous protein by the antibody, right?

Matthew- My cells are very healthy but I didn't actually knew that apoptotic cells are more florecencent that healthy cells

Alsadair- I am using 10% Normal goat serum and I am using the recomended conentration of the primary and secondary antibodies but it wouldn't hurt to go lower on the primary and secondary antibody concentrations. Thanks a lot for your response

Kazuo - Thats a good idea I will try that. I transfected HEK 293 with the expression vector with the desired cDNA cloned and I am trying to look for localization of my protein usind IF. I have seen expression of the protein via western blot. Thanks a lot for your rsponse

Hi Priya,

you already got a lot of good answers, so I just like to add a small advice. I had some similar problems recently and could piont the problem to the antibody. My normal staining protocol (Antibodies diluted in 5%BSA in PBS with 0,1% TritonX100), didn't work for the primary antibody. Thanks to a collegue of mine I tried a protocol without any blocking solution and it worked pretty fine. Here the protocol, if you want to try it out:

wash in PBS 5 min - incubate 90 min up to 3 hrs with primary antibody diluted in PBS + 0,2%TritonX100 (roomtemperature) - wash 2 times for about 5 min in PBS with 0,1% Tween20 - incubate 70 min with secondary antibody diluted in PBS + 0,2%TritonX100 (roomtemperature, dark) - wash 2 times for about 5 min in PBS with 0,1% Tween20 - counterstain with DAPI 5 to 10 min (depends on your experience with your DAPI) - wash 5 min in PBS - wash 1-2 min in H2O - mount (if necessary for you mounting medium dry before mount).

Maybe it helps. Last comment: as Alasdair already said, its difficult to judge without pictures, but apon aging DAPI sometimes makes a fluorecence shift and you will see it as background in the green or even the orange spectra (depends also on your filtertyp). To check this out, just have a look on your staining before DAPI-counterstaining.

Hi Anja,

Thanks I have read about some protocols which do not use blocking and I was a little skeptical not using the block as I already have background issues and I didn't knew of anyone who ever did without the block. But now after your response I will try to do without the blocking and see if the problem gets solved. Once again thanks a lot for sharing your protocol.

You're welcome!

Hi Priya,

I am right now using HEK293 cells for my IF studies transfecting gene of my interest. You see background for all above mentioned reasons.

Now no primary control..how does that look? Do you really see lot of background already with your secondary ?

may be you can change the secondary and see if you still get the same background. The higher you go towards far red in wavelength, you tend to see more of it. just to rule out you could try different colour.

During the procedure and your fixation protocol does contribute to dryness and background so what fixation? Methanol or PFA?

Blocking is a new point. Even I am not aware of skipping blocking but trying is good. Washing with detergent 0.1% Triton X but yes it will not give you a profile if your protein localizes to mebrane as detergents will rupture the memb. and shpw predominant cytoplasmic or nuclear staining wherever your protein goes.

So if you want to do no-permeabilization then you need to avoid detergents and use only PBS for everything. Increase the wash timings and no. It works fine for me.

Yes you can also reduce the amount of plasmid you are using for transfection. Sometimes over expression can cause high signal and background is seen even though cells are not transfected in the same field.

Good luck.

Hi Kapita,

Thanks for your response . My no primary control doesn't show any stainting so my secondary antibody seems to be fine. But I like the idea of transfecting less of the vector and also I am looking at a membrane protein and I am aware that detergent treatment will distrupt the membrane but the problem is my primary antibody binds to theN terminal of the membrane protein which is cytoplasmic so I am worried if I don't use any detergent how will the antibody have acess to the cytoplasmic N terminal.

HI, this is what I am doing exactly!

membrane protein but I am able to pick up my signal with a c-term AB where both C and N term of my protein are cytoplasmic. Well what happens is with whatever percent reagent you fix the cells, you surely pick up the cytoplasmic exposed epitopes I guess. I recently tried live cell and even there my Ab picks up signal which simply means that in 3D topology you still have some exposure. Actually membrane protein detection is little tricky until you have a fusion construct as the treatments of washes and fixation still make membrane porous to an extent.

Very mild detergents which do not solubilize membrane but make a hole are available , just in case. Such as digitonin, saponin, tween 20/80 etc.

Hi Bastian,

Thanks a lot for sharing your protocol. The protein I am looking at is membrane protein with 10-12 transmembrane segments. Also according to the western blot ,on which my antibody works well, HEK 293 do not express my protein. I am trying to look for different antibody that will work but giving my best shot at the current antibody that I have before I discard it for IF

Priya

Washing step should be important, otherwise put one drop of BSA or Goat serum mount with cover-slip see the auto-fluorescence.

The newest and greatest blocking agent is a marine blocking agent (MAXblock). I have found this to be the absolute best blocking agent. I read about it in a methods paper. Here is a link in case you want to check it out!

http://escholarship.umassmed.edu/cgi/viewcontent.cgi?article=2596&context=gsbs_sp

Good Luck!

Hi Andrea,

I had never heard of this blocking solution seems like it would be worth trying out. Thanks a lot for sharing the publication

Priya

Dear Priya,

my experience - I work for a company that develops diagnostic tests on the basis of HEK293 that express either of a multitude of autoantigens for IF serology in human samples - is that in most cases commercial antibodies do not work very well for immunofluorescence. Most of them are raised with peptides and are only controlled for reactivity against either the peptide or the denatured protein (westernblot). The idea of the IF is to use a protein in a near authentic environment, i.e. folded, modified and connected to all sorts of other biomolecules. So, if you have the chance of using another antibody try that.

Blocking is almost never an issue. You can go as simple as PBS (all other buffer subtances work worse in our experiments) + 0.2-0.5% Tween 20. If you like add BSA or goat serum (but that is what you do anyway).

Another issue is the fixation: one of the mildest fixations is done with pure acetone. You get rid of all membranes BUT you might get unwanted staining (with sera not so much with antibodies) and the location of target might get changed because it can roam around. However, it should be a good way to find out if your antibody binds to something. With formalin (be sure to get methanol/acetic acid-free formalin!!!) you keep the cells closed and really fix all proteins in their places BUT you might end up destroying the epitopes that your antibody binds to due to modification of amino acid side chains. And you have to find a means to let the antibody penetrate (detergents in different concentrations - Fab fragments are much better for this because they are much smaller and can enter better).

Lars

Dear Lars,

Thanks a lot for yuor response. I am planning on using a different antibody. But I was kind of wondering if there would be a way to dentaure the protein for IF like treatment with SDS ?

priya

SDS (or other harsh detergents) will rather detach your cells than do any good. And you would certainly loose any information on the location.

You could use your GFP-tagged variant and another secondary conjugate (red) for a double staining and optimization of your protocol. If that gives you any clue on the applicability of the ab you can turn back to the untagged variant.

Hi Priya,

even after having cross-read all the previous answers and comments, I'm still not quite clear about your problem. "Background staining of untransfected cells" -

... How do you know they are untransfected? 293 are so transfectable, a good protocol will give you very close to 100% transfection rate.

... As a negative control, did you stain a non-transfected sample? What about possible endogenous epitopes? Make sure 293 is definitely a null background for your protein, and also for proteins with similar N termini.

... What about expression levels from your vector? When you massively overexpress a membrane protein, you may overstrain the cell's protein processing machinery, and the half-folded protein will basically get stuck in the ER/Golgi compartment. Very little of it will actually reach the plasma membrane, and you may be fooled into believing this is "background". In this case, using a weaker, cellular promoter (e.g. UbiquitinC or EF1-alpha), instead of the usual, strong viral promoters (e.g. CMV) will help - more than using less plasmid.

... If this is not an option, try a different host cell type. Cell types differ vastly in their ability to process high levels of a membrane protein they would not normally express. Some neuronal ion channels express properly only in - neurons.

... "Clean" plasma membrane stainings are not always easy to see. 293 cells have surprising morphologies, once you see the actual cell margins. Make sure you're looking in the right place. Plating the cells at low density, and co-staining with a membrane marker, or with an actin label (e.g. Phalloidin), will help.

I agree with previous comments, about blocking not being an issue in IF. It made sense back in the childhood days of secondary ab's, to block with serum (= non-labeled IgG) of the species in which the secondary was raised. Hence the goat serum, as most secondaries were raised in goat. It served to saturate possible IgG binding sites in the sample with non-labeled IgG, preventing labeled, nonspecific IgG from binding. Most secondaries of today are of very good quality, so in most cases this is no longer necessary.

Also, it no longer hurts when people use goat serum for blocking, and donkey or chicken secondaries ;-)

Hope this heps,

Alex