Hi Dayan, I think Cecile is right, you got it a bit mixed up with goats and donkeys... ;-)  So, there are essentially no primaries in donkeys yet, or maybe very few. But since there are more and more primaries made in goat nowadays, the Companies are shifting to making secondaries in donkey. Possibly you also have the advantage of a higher yield in the bigger animal. Anyway both secondaries are regularly used, there is no real advantage of one over the other.

In any case if there are any primaries in donkey, they are useful for multiplexing. If you need to detect more than two proteins at the same time, mouse and rabbit are not sufficient, so you can use a chicken or a guinea pig, or a sheep, or a goat, or a donkey, etc. primary for the third and/or fourth protein (typically modern LSMs have 4 lasers: UV (405 nm), blue (488 nm), green (540-560 nm) and red (630-640 nm)) allowing the detection of 4 dyes in the same sample.

So, for your case: you have validated mouse and rabbit primary Abs, just stick to them and choose goat or donkey anti-mouse and anti-rabbit secondaries. Theoretically it is best to have the secondaries in the same species, this because you will use for blocking the serum from that species. This should reduce the background from the secondaries by blocking possible non specific binding sites for the goat or donkey Abs. In practice, most of the non specific signal normally comes from the primary Abs, background from secondary Abs is rare (anyway try a slice with secondary only every time you have a new batch or you change staining conditions), and that would be one case when you try to use higher dilution of the secondaries, although higher than 1:1000 in IF you normally don't go, mostly for confocal imaging, or you'll have too low signal to image anything (keep in mind that in IHC you have an enzymatic amplification, in IF you don't, the signal comes only from what is bound). As Cecile says, typical Alexa conjugated secondary dilution is 1:400 - 1:500. But keep in mind that Alexa conjugated Abs are VERY batch dependent, some work very well at high dilution, some just work very bad and you can only try to concentrate or buy a different batch (essentially they are too weak, either because of too little conjugation or due to inactivation of the Ab during the chemical reaction to conjugate them).

Second point, the Triton story. Triton has generally very little effect on protein structure and protein-protein interactions because it is a nonionic detergent while interactions mostly work on ionic interactions and proteins expose a ionic surface to remain soluble in water, so the detergent is kept out. But Triton is essential in slices to keep the membranes open. Since you fix the tissue before staining and since Triton does not dissolve the membranes but produces small mycelles, they remain trapped in the tissue due to the fixation of the extracellular matrix, so removing Triton from any buffer will produce rejoining and rearrangement of the mycelles, creating large membranes sheets that interfere with the penetration of the Abs. But normally a low concentration is sufficient, between 0.2 and 0.3% in all buffers, from blocking to washes. Besides, it also reduces surface tension during the staining procedure, making easier and less stressful for the slices the buffer changes.  For certain Abs a milder nonionic detergent increases staining efficiency, Tween. This you should determine empirically for your specific application, in case you are not happy with the staining using Triton.

A better trick when staining thick slices (so from 20 um on) to increase the penetration of the Abs through the fixed extracellular matrix deep into the tissue is to also add 1-2% DMSO to the blocking solution (in which one also normally dilutes the Abs). This is a polar solvent that interferes with both polar and non polar interactions, but at this low concentration is not disruptive for Abs interactions, although it helps to create holes in the fixed matrix and to expose epitopes.

Last point concerning the buffer that can be very important is its pH. Typically people use PBS at pH 7.4. If this works fine, no problem, but often phosphate buffers can form precipitates with the Ca2+ in the tissue (mostly in brain!), trapping Abs and producing the classical spotted staining. Then better to use Tris based buffers such as TBS. Abs work best at pH 8, although this is not always good for the tissue or for some antigens. Again, it should be tried empirically which pH, 7.4  or 8.0, works better.

Then of course there are all the possibilities of incubation at 4 °C, RT or 37 °C for short time, O/N or several days. When a staining does not work in standard conditions you should go for the whole panel of possibilities. For confocal microscopy you need to maximize the staining efficiency, otherwise the quick quenching due to the strong laser illumination and the high loss of fluorescence signal due to the confocal approach will leave you with background or no signal...

Good luck.

IHC is an art,  typically you should titrate you antibodies for your tissues. How is you brain fixed? which species are your brain from? - usually people use donkey or goat for convenience, it should not make any difference. Triton as well is an art, we used 1% triton for blocking and primary incubation and 0.3% with the secondary antibody, again this depends on how you fixed your tissue. It's OK to dilute your ab when you use DAB, but for fluorescence, we always used more concentrated ab (1:200-1:400), again you need to titrate this for your own experiments, and have a good balance signal/autofluorecence. Check this paper for mouse brain IHC, Radiat Res. 2008 Feb;169(2):149-61. I unfortunately do not have the paper on file readily available

Hi Dayan, I think Cecile is right, you got it a bit mixed up with goats and donkeys... ;-)  So, there are essentially no primaries in donkeys yet, or maybe very few. But since there are more and more primaries made in goat nowadays, the Companies are shifting to making secondaries in donkey. Possibly you also have the advantage of a higher yield in the bigger animal. Anyway both secondaries are regularly used, there is no real advantage of one over the other.

In any case if there are any primaries in donkey, they are useful for multiplexing. If you need to detect more than two proteins at the same time, mouse and rabbit are not sufficient, so you can use a chicken or a guinea pig, or a sheep, or a goat, or a donkey, etc. primary for the third and/or fourth protein (typically modern LSMs have 4 lasers: UV (405 nm), blue (488 nm), green (540-560 nm) and red (630-640 nm)) allowing the detection of 4 dyes in the same sample.

So, for your case: you have validated mouse and rabbit primary Abs, just stick to them and choose goat or donkey anti-mouse and anti-rabbit secondaries. Theoretically it is best to have the secondaries in the same species, this because you will use for blocking the serum from that species. This should reduce the background from the secondaries by blocking possible non specific binding sites for the goat or donkey Abs. In practice, most of the non specific signal normally comes from the primary Abs, background from secondary Abs is rare (anyway try a slice with secondary only every time you have a new batch or you change staining conditions), and that would be one case when you try to use higher dilution of the secondaries, although higher than 1:1000 in IF you normally don't go, mostly for confocal imaging, or you'll have too low signal to image anything (keep in mind that in IHC you have an enzymatic amplification, in IF you don't, the signal comes only from what is bound). As Cecile says, typical Alexa conjugated secondary dilution is 1:400 - 1:500. But keep in mind that Alexa conjugated Abs are VERY batch dependent, some work very well at high dilution, some just work very bad and you can only try to concentrate or buy a different batch (essentially they are too weak, either because of too little conjugation or due to inactivation of the Ab during the chemical reaction to conjugate them).

Second point, the Triton story. Triton has generally very little effect on protein structure and protein-protein interactions because it is a nonionic detergent while interactions mostly work on ionic interactions and proteins expose a ionic surface to remain soluble in water, so the detergent is kept out. But Triton is essential in slices to keep the membranes open. Since you fix the tissue before staining and since Triton does not dissolve the membranes but produces small mycelles, they remain trapped in the tissue due to the fixation of the extracellular matrix, so removing Triton from any buffer will produce rejoining and rearrangement of the mycelles, creating large membranes sheets that interfere with the penetration of the Abs. But normally a low concentration is sufficient, between 0.2 and 0.3% in all buffers, from blocking to washes. Besides, it also reduces surface tension during the staining procedure, making easier and less stressful for the slices the buffer changes.  For certain Abs a milder nonionic detergent increases staining efficiency, Tween. This you should determine empirically for your specific application, in case you are not happy with the staining using Triton.

A better trick when staining thick slices (so from 20 um on) to increase the penetration of the Abs through the fixed extracellular matrix deep into the tissue is to also add 1-2% DMSO to the blocking solution (in which one also normally dilutes the Abs). This is a polar solvent that interferes with both polar and non polar interactions, but at this low concentration is not disruptive for Abs interactions, although it helps to create holes in the fixed matrix and to expose epitopes.

Last point concerning the buffer that can be very important is its pH. Typically people use PBS at pH 7.4. If this works fine, no problem, but often phosphate buffers can form precipitates with the Ca2+ in the tissue (mostly in brain!), trapping Abs and producing the classical spotted staining. Then better to use Tris based buffers such as TBS. Abs work best at pH 8, although this is not always good for the tissue or for some antigens. Again, it should be tried empirically which pH, 7.4  or 8.0, works better.

Then of course there are all the possibilities of incubation at 4 °C, RT or 37 °C for short time, O/N or several days. When a staining does not work in standard conditions you should go for the whole panel of possibilities. For confocal microscopy you need to maximize the staining efficiency, otherwise the quick quenching due to the strong laser illumination and the high loss of fluorescence signal due to the confocal approach will leave you with background or no signal...

Good luck.

Wow!!!!!! thanks for the thorough responses. So I fix section in paraformaldehyde. Rat brain sections sliced at 30um. I'll take all of these into consideration when troubleshooting my protocol. Again, thanks so much.

To piggyback off of Ceclie and Peitro's answers, IHC using immunofluorescence is truly an art form ( not without the headaches!)

Whenever I've done double, or triple labeling with fluorescence, I use a 0.5% concentration of Triton for the Primary Antibodies, and 0.1% for the Secondary. Do you use the 0.1% concentration for both primary and secondary? If you can, play around with the concentrations.

With secondaries, it's pretty standard to have your concentrations be at pretty high dilutions ranging from 1:250-1:500. What's the dilution you're using for your primaries- and what specifically are you staining for? I've found its usually the primary dilutions that are an issue, or that one will work with DAB and not with fluorescence. It's all finicky, until you get that magic concentration that the science gods graced you with.

www.ihcworld.com has been a go to for me whenever I have these issues, hope that might help out as well.

good luck and my fingers are crossed for you! Feel free to inbox me as well.

-Griffin