Sometime, under florecense microscopt, the cell show the color, however, it is not the fluorescence you need. The real positive signal are stronger. 

You can dilute more when using your second anitoby. Can the real signal is not in every cell. You will find the difference between the positive signal and negative signal. 

i will dilute the secondary and see how it looks

Try serial dilution (1:2 or 1:10) until the one you barely see fluorescene in negative control the use that concentration for your samples.

Hello Jaya,

To be safe, check the literature available on the cell line you are using to make sure your cells do not express any Fc receptor that could bind to your secondary antibody.

Good luck

Cecile

hi jaya,

I am agreeing with all previous mentioned answers.

What about washing? how many times did you wash after the secondary Ab?

I washed three to five washes with 5 min interval

What is your dilution? How are you blocking and permeabilizing your cells? What is your detection methods: widefield? confocal?

For AF488 secondaries I regularly use a dilution of 1:1000 and block with 5% BSA in PBS+0.25% Triton X-100. Washing is important, but your dilution, block and perm methods will also have a significant effect.

fixing in formaldehyde, permeablizing in 0.1% triton X100 and blocking with 5% goat serum. Sec 1in 1000 for 1 hr and washing is 3 time with 5 min interval and I dont block with BSA. The images are taken in widefield.

Do you have another color of secondary you could use to confirm it isn't just specific to that batch of antibody?

Yes I do and I will use it and check for the result.

I agree with the Jordan. I also used BSA for blocking, dilution 1:2000-1:3000 (both worked). Washing 3 times with 5 minutes interval. Sometimes to reduce the non specific binding, you can try to reduce the incubation time of secondary to 30-45 minutes or increase your washing time.

U r using sec Ab from which company? Sometimes batch to batch variation is there. U can call them and ask to replace for free, if you have tried all the alternatives.

Good luck

Well, you seem to be doing everything right. Here are a few long shots:

1. Blocking: I am assuming your secondary is goat (since you are using goat serum to block).  Are you just removing the blocking solution and replacing it by the primary (no ringing in between)?  Are you adding some goat serum in the primary Ab dilution (this can keep the blocking effect).

2. Rinsing: are you rinsing the cells in buffer before fixation (or serum free media).  The secondary could react with some serum components.

3. Drying: Are you sure the cells don't dry out during the process.

4. Substrate: Is you substrate special and could it bind the AB.

5. PPTion: Is you secondary filtered/centrifuged.  Sometimes the secondary can aggregate in small fluorescent particulates that bind to the cells.

6. Are your cells from the same specie as you primary (e.g. mouse cell if the secondary in a goat anti-mouse)?  Then that would explain it.

Next time, you should add more details to your question so we don't have to guess at the context ;-)

You may want to double check that you are using the correct serum for blocking in both the primary and secondary antibody stage. These should coincide with the species of animal in which the antibodies were made.

There are also Protein-free blocking agents made by companies ( Millipore has one I think ) that are claimed to provide for less background staining.

I would like to comment on Maureen's comment.  You DO NOT want to block with the serum of the animal species the made the primary antibody.  Use only the serum from the species that made the SECOND antibody otherwise you are guarantied a high background.  If you coat the target cells with, let's say, mouse serum to bing to the nonspecific sites and then treat it with goat-anti-mouse labelled antibodies, you will light up all the antibodies from the mouse serum that adhered non-specefically to the target.

In response to Nicolas l'Heureux, I meant to block with the first antibody species with the Primary Ab and second antibody species with secondary Ab.

Thank you all for your suggestions and sorry for not giving enough details. Iam using human cervical cancer cell to do immunofluorescence and Iam probing for two proteins MMP20 using rabbit poly clonal and DSPP using mouse monoclonal.

The protocol that I follow is

1. wash the medium and wash the cells twice with pbs

2. Fix with 4% formaldehyde for 10 min, wash twice with pbs

3. add 0.1% triton X100 for 5 min and wash twice or thrice with pbs

4. Block with 5% goat serum for 1 hr and add the primary ab (in goat serum) two hr at RT or 4deg overnight.

5. wash the cells thrice with pbs at 10 min interval and add sec  alexa 488 (goat anti mouse) and alexa 555 (goat anti rabbit).

6.wash thrice with pbs at 10 min interval ( sometimes I keep in a rocking platform at very low speed).

As a negative control, I do the same with Ig G instead of the primary ab. I problem Iam facing is in the negative control both rabbit and mouse the nucleus of the cell is lighting up but not the cytoplasm as if its a nuclear stain.

Trouble shooting.

a. Instead of 5% goat serum, I used fab blocking and did the experiment only with Ig G. same result.

b. Diluted the primary at 1:200, 400 and 600. Same result

c. increased the dilution of secondary. 1: 1000, 1500 and 2000-  same result

d. Purchased a new Ig G- same result

e. Fixed the cells in PFA instead of formaldehyde- same result

f. cells are not autofluorescent

g. repeated the expt with the secondary from other lab- same result

g. only primary or only secondary did not give any signal at any of the above mentioned dilutions

But when the primary and secondary are added the fluorescence is seen and clearly in both red and green its only the nucleus is stained. Someone even asked me if Ihad switched the color for DAPI, as it was so prominent.

Any suggestions are welcome. SInce Iam not able to fix the problem in the negative control, Ihave stopped doing any expt for the past two weeks and just concentrating on this,but not able to find where the problem is. Please help

Jaya

You are performing the immuno on human cells correct? I would double check with Alexa that the IgG you are ordering has limited cross-reactivity with human cells. Their 488 goat anti-mouse polyclonal has limited cross-reactivity; not sure about the monoclonal you are using. But if the antibody has not be pre-adsorbed against human, you will have cross-reactivity with endogenous IgG in your human cells.

You can also increase the % of goat serum in the blocking solution. I typically use 5% as you do, but I have gone as high as 10% with amazing results - usually if I do that, I increase secondary incubation time, but given that your secondary is binding nonspecifically, I would not advise that.

You can also try diluting further the secondary, if you haven't already.

Also, when you do secondary alone, is the fluorescence localized to the nucleus, like you were seeing before?

Thank you Caitlin. I can increase the conc of the blocking as 10 %. when I use the secondary alone at any concentration that I used I dont see any signal. Its only with the primary Iam seeing it.

Jaya,

In your initial comment you said that "Its the secondary alone that is giving very high fluorescence. " and now you say that "g. only primary or only secondary did not give any signal at any of the above mentioned dilutions".  These seem like contradictory informations. It does seem from your later comments that  the secondary alone does NOT give a signal. I just want to make sure I understand. 

You say that "I problem Iam facing is in the negative control both rabbit and mouse the nucleus of the cell is lighting up but not the cytoplasm as if its a nuclear stain."

You also say that "when the primary and secondary are added the fluorescence is seen and clearly in both red and green its only the nucleus is stained. "  Are you saying that your negative control (non-immune IgG) gives the same staining as when you use the specific IgG?  In your negative control, do you mix some noontime IgG from mouse and rabbit.

Also, When you dilute the secondary ("c. increased the dilution of secondary. 1: 1000, 1500 and 2000- same result"), do you see a decrease in the signal?

When you used secondary antibodies from the lab next door, were they from a different provider or just a different batch?

Im sorry for the confusion. Its the negative control showing fluorescence. When tested individually, they do  not show any fluorescence, but when together the nucleus is stained. I originally thought that it was the DAPI giving some fluorescence, so eliminated it all together. Even after that I see the fluorescence in the cells. The negative control is mouse and rabbit and Its done individually. I take care to see that there is no cross contamination as well. 

When I increase the dilution of the secondary, there is no decrease in the signal. So Iam thinking that the Ig G is binding to the cells. The secondary that I got from the other lab is from a different company.  Now Im have fixed the cells in PFA instead of formaldehyde that I was using earlier. I have to view the result. I will post it if there is any difference.

Hi Jaya:

Try to not using seurm as your blocking agent at all. Instead you can use freshly prepared Sodium Borohydride  (5mg/ml) for 10-15 min at RT or 3%gelatin-PBS at 37 C for an hour and rinse them with PBS warmed to 37 C. Serum can bind to the cells non-specifically and be recognized by the secondaries. Good luck.

Thank you. I will try it

Hi Jaya, I'm experiencing similar problem as you. Therefore, i'm curious if you manage to find a solution and the crux of the issue. 

Breast cells on coverslips, Rabbit Primary, Goat anti-Rab Sec (488)

I'm using 4%PFA-10m , Triton 0.5% 5m, 5%FBS/BSA block 30m, (3rinse2min each) 1Hr pri @ RT,  (3rinse2min each), 30m Sec Ab @RT  (1:400 aka 50ug/ml), (3rinse2min each), DAPI 3min ...

I'm seeing cytoplasmic/some nuclear in my secondary-only control coverslips likely false staining..

Dear Jaya,

I read all the comments so far and what I think is totally different than previous comments. I think there is nothing wrong with your protocol. Instead I think the problem is with your cells. Your cells are not very healthy, possibly very depolarized membrane and they have very high permeability. Although you are fixing them your cells already have large pores on them. When the cells are unhealthy like this it does not matter what you do/try you will always get non-specific labelling. You can try incubation without any permeabilization with Triton-x 100 and you will find out that there will be exactly the same amount of nuclear/cytoplasmic labelling. Nucleus is usually very protected and the pores on the nuclear membrane (nuclear pores) are only open non-specifically when the cells are extremely depolarized causing excessive cellular stress.  I am an electrophysiologist and I regularly do IHC on cells after recording their membrane potential (see my previous papers in RG). I know that the only time you get non-specific labelling with your secondary antibodies is when your cells' membrane is very depolarized (membarne potential is more positive than -30 mV). At that resting potential many sodium and calcium channels open and cells are almost certainly set to die and the membrane integrity is lost and the pores open up all over.

My advise is try your secondaries in healthy cells and you will notice that there is nothing wrong with your protocol. If you happen to know an electrophysiologist around ask them to check the membrane potential of your cells.

best wishes,

Refik

Hello everyone,

I had the same problem and I found the same things which were described by Dr. Refik Kanjhan: if you compare the background of secondary antibodies only between healthy and dying cells, you´ll find the huge difference. So, thank you so  much Dr. Refik Kanjhan for the explanation about the depolarized membranes. The only problem is how to interpret such data?) 

Thank you

Dear Jaya,

Did you find solution to your problem? I am in similar situation right now. How did you resolve the problem. Any tips and tricks are welcome.

First see the cells are healthy because dying cells show fluorescence.

Second fix the cells with 4% Para formaldehyde, not with formalin or depending upon the antibody used, methanol can be used. Hope this helps

Hi Jaya,

When tissue samples fixed with an aldehyde fixative such as formalin or paraformaldehyde and immunofluorescence (IF) is the detection method, you need to block free aldehyde groups that are generated during the aldehyde-driven crosslinking (fixing).

The most effective strategy to eliminate these reactive groups (which will inevitably bind your primary and/or secondary Abs) is to include 0.3M glycine in the blocking buffer. Also, remember that the serum you are using in the blocking MUST be from the same animal species of your secondary Abs.

Also, remember that adding 0.3 M glycine will lower the pH, so make sure to adjust the pH after adding the glycine to your blocking.

Another important rule is to use secondary antibodies as F(ab)2 fractions (NOT whole IgGs) from a large animal from which companies can harvest large batches of serum, and can afford producing cross-species preabsorbed secondary antibodies.

I personally like to use the F(ab)2 fluoroconjugated Abs from Jackson Immunoresearch Labs, and select their ML versions that are preabsorbed on multiple species.

Their secondary F(ab)2 fluoroconjugated ML Abs from Donkey are great and superclean, and they have normal (non-immunized) donkey serum to be used in the blocking.

Good lack!

Vin

It is simple. Alexa488 is a flourophore and suppose to give you fluorescence with appropriate excitation light. If you want you can also purchase primary antibodu conjugate with Alexa then you donot need secondary antibody.

Thank you

Hi Jaya!!

Please, what perfect way to solve this problem. Bcos I am currently in the same situation you were 3years ago. I will be glad if u can share how you solved the problem then. Thanks