Double immunofluorescence labeling when one fluorophore is amplified by TSA

I am trying to use immunofluorescence to label two proteins when one of the two fluorophores is amplified using a TSA kit (Invitrogen). Unfortunately, the second fluorophore (not amplified) is very dim compared to when it is imaged alone. I have applied the second fluorophore both after the TSA procedure and before and it doesn't seem to matter. Also, I block for at least an hour between protocols. Any suggestions?


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  • Viktor Kharazia · Merritt College
    interesting problem, are these cells or tissues? My guess would be that proteins of interest are really close and some sort of FRET is occurring
  • Viktor Kharazia · Merritt College
    I would try to reverse colors (expensive...)
  • Anna Costagliola · University of Naples Federico II
    I would suggest reading the following paper:
    Triple immunofluorescence staining with antibodies raised in the same species to study the complex innervation pattern of intrapulmonary chemoreceptors.
    Brouns I, Van Nassauw L, Van Genechten J, Majewski M, Scheuermann DW, Timmermans JP, Adriaensen D.
    J Histochem Cytochem. 2002 Apr;50(4):575-82.
    also reversing fluorophores might help
  • James Meabon · VA Puget Sound Health Care System
    Wow all kinds of things could be going south...
    Let's start from the top and try to cover the most basic ones...
    However, if you want the SPOILER SKIP TO PART 4...

    1) Fixation:
    You can try different permeabilization and fixation techniques. It'll save you a lot of time by trying a wide array at first to see which ones provide the best staining. The ability of an antibody to find and bind its antigen is often very dependent on these sorts of things. The problem often stems from a researcher finding one way that works for something and carrying on with that technique ever after. Different options include methanol fixation, acetone fixation, 4% PFA, transcardial perfusion followed, the list goes on.

    2) Secondary choices:
    Some primary antibodies are finicky. Some actually care if you try to target them with an fc-fragment or a full-length Ig secondary. You may want to start with trying an fc-fragment (or full-length if you already employ a fc-fragment as your secondary). These generally have better penetration and wash off better leaving a lower background.
    You also want to have a secondary that is best matched to your laser (obviously) but that has the best extinction coefficient (E)... meaning that if your secondary is an exact match for your laser but has poor E then you'll want to choose one with the highest E you can get with in a "reasonable" nm deviation of the laser you want to use. For Reds, Cy3 and 555 are generally very reliable.
    That may fix your problem. Of course it may not. You may simply be having problems with your secondary antibody finding your primary. In this case you can also try to see if you can conjugate your fluorophores to your primaries before use to ensure maximal signal. One of the problems with this is that most antibodies are supplied in buffers/formulations that contain primary amines (eg Tris or BSA) which compete with the desired reaction. You can try to do a microdialysis-- but it's usually low yield and therefore proportionately more expensive. If you don't have luck with the procedure/kit to do this (several are offered by Molecular Probes) or you just want to save time/money you can often contact the supplier of your primary to do it for you or atleast supply it in a formulation compatible with the conjugation reaction.

    3) Mounting:
    Your secondaries can dissociate from the primary over time and a fluorescence change over time can be seen. Usually there are three common solutions to this. One, do a post-fix after all the washing steps and before mounting. Two, don't use a liquid mounting solution like vectashield or 50% glycerol. Use a hard mount like ProLong Gold antifade (stuff is awesome! and no, I'm not a sales rep). Or three, image it immediately after mounting.

    4) SCAN ORDER:
    Since the second fluorophore is "dead" by the time you image it, it sounds like you are bleaching it by virtue of imaging your other fluorophore first. If I had to bet money, I would bet this is what is going on in your case...
  • Pamela David Gerecht · University of Colorado
    You wrote, "Unfortunately, the second fluorophore (not amplified) is very dim compared to when it is imaged alone." Did you mean its dim when it is the only Ab in the cell or when only this fluorophore is imaged in the presence of the other? If dim when used alone, then you need to try some of the things suggested by James above (1,2,3). If this antibody has strong fluorescence when used in the absence of the other TSA amplified one, then you either have FRET or one Ab blocking the other.
  • Clark Lindgren · Grinnell College
    Thanks everyone. I don't believe the problem is FRET or fading since it doesn't matter what order I image the antibodies. The ONLY time I have the problem is when I am using TSA to visualize the other antibody so I think somethin in the TSA procedure is interfering. Has anyone ever heard of this happening?
  • Brian Lin · Tufts University
    If you are expecting co-localization of the TSA'ed antibody and the second one, yes. If they should be completely non-co-localized, TSA should not affect your second ab staining.

    Our lab does a ridiculous amount of IHC, especially TSA, and its our belief that the deposition of the tyramide severely blocks epitopes, to the point that we use TSA to effectively do double or triple staining with antibodies raised in the same species.

    Have you tried to first stain the weaker antibody, and then TSA? This would effectively negate the blocking effect of TSA.

    How are you amplifying the weaker antibody? are you directly conjugating to a fluor? If reversing the order you do the staining does not help, I would highly recommend using another enhancement procedure.
    Let's say for ease of typing that your weaker antibody is a rabbit ab. You would use a fitc or texas red anti-rabbit (jackson immuno has these for ~80 bucks). Then use a rabbit anti-fitc/TR. (Again, jackson). Finally, do a round of fitc/TR anti-rabbit. This is not even close to TSA amplification levels, but there are several advantages.
    1) It does not use the biotin system so it won't interfere if you are using the biotin-tsa system.
    2) you can chain the system several times. We've done the enhancement 3 times in a row for excellent results.
    3) It's tunable, as you can stop the enhancement every other step depending on how bright you want it.

    Are you using the biotin-tyramide and then SA-fluor? Or are you using a directly conjugated tyramide-fluor?
  • Clark Lindgren · Grinnell College
    Yes, we are expecting colocalization; however, we have reversed the order and still had the same problem. I will try your suggestion and use another amplification procedure. We have been using the biotin-tyramide/ SA-Fluor procedure.
  • James Meabon · VA Puget Sound Health Care System
    Brian, could you explain how you use it to do double and triple labeling with same specie antibodies (like rabbit anti's, for example). I'm really curious; this sounds very neat.
  • Brian Lin · Tufts University
    Clark Lindgren, that's unfortunate, of course the easy solution wouldn't work :( Other than the fitc and TR enhancements, if that is not enough, you could get a second TSA kit that uses a directly conjugated tyr, and do two step staining, and just inactivate the HRP after the first development. Of course that's another 400 bucks or so for another kit, but that is an alternative.

    James Meabon, to use same species antibodies takes several days, more for each additional antibody.
    You basically put on the worst/weakest signal antibody first, and develop using TSA to completion. The deposited tyramide is able to mask the primary antibody from detection.
    If you are only doing two antibodies, you can then simply put on your second one and detect using conventional methods. If you need TSA for both antibodies, you must use different tyramides and inactivate the previously deposited HRP by washing with h2O2 in pbs. This works best if you use two fluor-tyrs, like cy3 or cy5 because you reduce the need to use biotin which would cross react the next time, and it decreases background in general.
    If you are doing more than 2 antibodies, it gets a bit dicey as the background amplification from TSA adds up since you have to do TSA for n-1 times, each time to mask the previous epitope. I've been able to do 4, although that took forever, and requires heavy-handed use of BSA-PBS wash which greatly decreases background. You also need as many different fluorescent tyramides. However, the results are pretty awesome once you get over the time investment for it.
  • James Meabon · VA Puget Sound Health Care System
    Interesting... I'll have to keep this is in mind. Thanks alot, Brian.
  • Brian Lin · Tufts University
    you are totally welcome! If you find some improvements or troubles, please let me know.
  • Eleonora Melzi · University of Glasgow
    Dear all,

    I have a similar problem. I'm trying to do a double staining on formaline fixed samples. I'm using two polyclonal rabbit and I amplify both of them using TSA of different colors (Alexa 488 and Alexa 594), but no matter which order I use the antibodies, I am losing the signal from the second used. When used alone both the antibodies perform well with TSA.

    I'm doing an indirect staining using secondary abs conjugated with HRP. I do the first staining using the first polyclonal, then I block hrp using H2O2, further block using normal rabbit serum followed by anti-rabbit fab fraction (all this to avoid cross-reaction). Then I perform the second staining using the other polyclonal.

    Any suggestion?
  • Brian Lin · Tufts University
    Are these two antibodies co-labeling the same cell/localization? The benefit that TSA lends by blocking epitopes could be blocking access to the second antigen.
    If not, then perhaps the H2O2 treatment is detrimental to your antibody epitope. This you can test by just pre-treating tissue with H2O2 and then staining a single antibody and see if your staining is not as good. If that is the case, the amount of H2O2 you use to inactivate HRP can be titrated to a minimum, and hope that is enough to preserve staining?
    I can't really think of any other ideas, unfortunately :(
  • Eleonora Melzi · University of Glasgow
    Thanks Brian,
    I am expecting colocalization just in a small part of the cells.
    But you might be right about the H2O2, I will try to reduce the incubation time with H2O2. I'm using it at 3% for 1 hour, I have tried for 10min but it does not deactivate HRP. Maybe half way could work.
    How long to you generally treat the sample to deactivate the HRP in a double TSA staining?
  • Brian Lin · Tufts University
    Interesting, are you dilution the H2O2 in diH2O or buffer? pH has an effect on how quickly HRP is inactivated, as more extreme pH, especially alkaline (I believe), the suicide catalysis is much quicker.
    For us, the only antigens that we use for TSA have been ones which actually withstand 3% H2O2 in MeOH, as it is a pretreatment before steaming to increase signal to noise.
    Our inactivation is 3% in water for 15 minutes. We do run the first reaction to completion, >15 minutes of tyramide incubation, which I believe masks quite a bit of the HRP activity itself.
    I have a feeling that 1 hour is quite long for 3% solution, generally 1 hour incbations are done at .3%.
    However, let me just list the three ways that people have shown HRP can be inactivated for IHC.
    1) 3% H2O2 in diH2O, 15 minutes and rinse. If you are using buffer, perhaps unbuffered water provides an accelerated inactivation? Just guessing now.
    2) .3% H2O2 in MeOH for 15-30 minutes. The lower percentage is supposed to allow for longer non-damaging incubation time and the MeOH accelerates HRP inactivation. I would definitely give this a shot as its generally used for tissue with lots of endogenous HRP, so perhaps it will be enough to clear it up for you.
    3) Finally, .3% H2O2 and .1% sodium azide for 10-15 minutes. This is a bit more hazardous, but as long as you aren't planning on drinking and eating off your dry powder measuring bench, you should be fine. The azide is a complementary way of inactivating HRP.

    Hopefully one of these helps your predicament!


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