Hi Steve, 

Here at Novus Biologicals, we use Triton solely during the permeablization step as it is a fairly harsh detergent - We permeabilize for 10 minutes.  Depending on how robust the cell type is, a shorter or longer time may be required for your experiment. However, in my experience it is best to choose an incubation time right in the middle to be sure that the cell membranes are not too permeabilized. After this step, we wash with 1X PBS three times and then switch to a .1% Tween-20 when making up the remainder of the buffers ( for example primary and secondary solutions).  I would recommend not using Triton for any step after initial permeabilization and then switching to Tween-20. 

Amelia 

Hi Steve, 

Here at Novus Biologicals, we use Triton solely during the permeablization step as it is a fairly harsh detergent - We permeabilize for 10 minutes.  Depending on how robust the cell type is, a shorter or longer time may be required for your experiment. However, in my experience it is best to choose an incubation time right in the middle to be sure that the cell membranes are not too permeabilized. After this step, we wash with 1X PBS three times and then switch to a .1% Tween-20 when making up the remainder of the buffers ( for example primary and secondary solutions).  I would recommend not using Triton for any step after initial permeabilization and then switching to Tween-20. 

Amelia 

I agree with Amelia. Be careful in using Triton. Triton is used to impermealize tissues. You have to use it with short times and before beginning immunohitochemistry. You can also use it during the preincubation step but at lower concentration if your preincubation step will be long.

If only it were easy enough to say that you should always do one thing or another for quality results from an IHC procedure.  See Journal of Histochemistry and Cytochemistry article by Goldenthal et al, 1985.  

The appropriate use of a permeabilization agent may depend on the antigen's location in the cell.  However, most labs use standard blocking and antibody solutions since customizing each to a particular antibody may be impractical.  We use 2% serum with 0.2% Triton for blocking, primary antibody solution, and secondary antibody solution with only a couple exceptions where we use Tween-20.  The discrepancy between our protocols and the ones mentioned above may be explained by several factors including fixation procedures, typical target antigens, antibody concentration, etc.  

There are a number of things to consider.

1) Time of fixation (longer fixation may require additional permeabilization)

2) Thickness and size of tissue (whole mount of mouse retina vs. 20 micron section)

3) Quality of antibody

4) As Justin mentioned above, the antigen's location within the cell.

For the vast majority of IHC staining  in our lab, we permeabilize before incubating with the primary.

However, in our lab it is not uncommon to use a 0.1% triton to dilute the primary antibody. This is true for thick sections and "difficult" antibodies. It may be not be the answer you want to hear, as you may need to troubleshoot. 

Short answer: stick to permeabilization before the primary. If staining fails (after incubation time and concentration troubleshooting) switch to 0.1% triton during primary incubation. If that fails, time for a new antibody!

We use an incubation buffer containing 5% BSA + 0.3% triton for both primary and secondary antibodies and so far have had no problems. In my experience the antigen retrieval method is usually the problem.

So it sounds like the general consensus here is that mixing triton with primary antibody should be limited when possible, but in instances where antigen access is a challenge it may be required.  Could anyone speculate on why triton can occasionally decrease primary labelling?  Is it denaturing the primary antibody and thus weakening the antigen recognition site?  Or does it interfere more directly with antigen-primary recognition?  Could this information be used somehow to predict which primary antibodies are more likely to be hindered by triton?

Dear steve,

I will try to give some explanations to your questions.

Could anyone speculate on why triton can occasionally decrease primary labelling? Is it denaturing the primary antibody and thus weakening the antigen recognition site? Or does it interfere more directly with antigen-primary recognition? 

As mentioned above, triton is used for antigen retrieval operation.  In this sense, triton acts as a detergent to let antigens more accessible. But triton is a strong detergent and its time action has to be limited (as shorter as possible). this is true when you use it prior incubation or during incubation with first antibody. To my knowledge, there is no interaction between triton and antibody but its action as detergent (notably during the period incubation which could be important) could deteriorate antigens.

Could this information be used somehow to predict which primary antibodies are more likely to be hindered by triton?

May be some antibodies could interact with triton. For this you have to read carefully the technical notice of the antibody you use. 

If you want to use triton during your incubation step, try to minimize its concentration. Alternatively to know which optimal dilution of triton you have to uses, you can test 4-5 dilutions during your incubation step and compare results.

For tissue slices, I agree with Alice.

For cultured cells, It is dependent on how you prepare your samples. if you use methanol or ethanol as the fixative, you do not need to permeabilize cells with Triton. If you use (para)formaldehyde to fix cells, using Triton  to premeabilize cells after fixation may let you get better results. However, you need try and try to get the best results. In my lab, we  use ice-cold 0.1% triton/PBS  for 10-15 min to premeabilize fixed cells. This procedure usually can help us to get good results when we label antigens located at the cytoplasm or in the nucleus. 

In my experience, antigen retrieval is the main issue during IHC. I have routinely used Triton-X100 at the concentration of 0.05-0.2% with no real problem. The concentration of Triton used is normally dependent on the strength of antigen-antibody interaction. In absence of Triton, I tend to see more background.

Triton is normally used only for cell permeabilization, and its concentration differs depending on the type of the cell you want to treat with antibodies. With human or mammalia cell cultures grown on the cover slips 0.1 % is perfect, but for some protists even 0.5 % will not work after paraformaldehyde fixation. With the first antibodies we normally use 0.1% Tween 20, which is more gentle. I would not add Triton at this step just because you may lose all the material, but not because it interferes with the antibody.

Same here as with Pankaj Sharma.  I work with formalin fixed paraffin embedded CNS tissue and use 0.1% Trition in the buffer for all steps.  Really cuts the background particularly in the myelin which tends to be "sticky".  

I would think the reduction in quality that Trition is giving in other instances might be due to doing its job too well!  I have seen that happen when over enthused folks use an antibody at too high concentration, over retrieve or warm the slides.  The antibody does link, but you have background ALL over the place and do not have contrast and good photography.  Perhaps the Trition over permeabilizes the cells and there is an increase of non-target binding.

Triton inhibits (unspecific) hydrophobic interactions. Therefore it's possible that this detergent can inhibit the interaction with a hydrophobic epitope. A pure hydrophobic epitope is rare but possible, but only for monoclonals. So it's always good to use either a set of monoclonals, recognizing the same antigen or an polyclonal antibody. The usual Triton concentration should be at 0.1% or below

To answer your second Q:  Since triton is a harsh detergent, and a potent permeablizer you can potentially remove the epitope from cells/tissue sections, especially if it a membrane bound antigen. Therefore reducing signal intensity. in predicting which Ab are hindered by tritonX, always investigate where the antigen is located in the cell, make sure sections/cells are optimally fixed for the Ab you are using,and finally run a optimization/validation steps before you put the Ab to use on your study tissue.  Optimization is very important in making sure the Ab stains the correct antigen, and the correct location on the cells.  There have been instances (personal experience, and published lit) where the wrong permebilizer was used and the Ab stained the wrong cellular compartment. I always trial  tha Ab with and w/o digestion step, as well as different epitope retreival procedures (HIER, EIER).  Different fixation maethods require different steps as well.  There is no one fit answer to this question.  Just make sure you have validated the Ab under YOUR conditions.

I'm using 40 um-thick free-floating mouse brain sections, and I recently discovered that my sodium citrate antigen retrieval method (80C for 30 min) greatly damages my tissue (rough surface) unless I put triton-X in the blocking buffer (0.3% is suggested by the ihc world protocol). Pretreating with 0.1% trition-X for 20 min is not sufficient to stop this damage from occurring. Not sure why.

Hi,

Could anyone tell me the time it takes for triton permeabilization to reverse in fixed cells? if found that its 24h for living cells but can imaging it takes longer for fixed cells.

thanks,

Hans.

0.1% triton X100 in blocking and primary buffers always work just fine on my OCT frozen 5um thin tissue sections. Some antigens are fixation sensitive, so try treating sections with blocking and primary before fixation if thats the case...Good luck!!!