Hi Catarina,

When I've imaged tagged nuclear proteins in BSF T. brucei in the past I've always used 2% PFA (15 minutes on ice) and permeabilised in 0.5% triton-X100 for 20 minutes. This works well in most cases. An alternative that I've used the one time this approach failed (for an antibody to a modified histone) was incubation in ice cold methanol for 10 minutes (no TX100 treatment required), though the cellular morphology is never as nice as with PFA fixation. One more thing, do you dry your cells onto ordinary slides or settle them onto poly-L-lysine coated slides? The latter often gives better results, at least in my hands.

Hi Sam,

Thanks for your reply. I always settle them onto multi-well poly-L-lysine coated slides. I'll try your suggestions in parallel with the protocol I've been using and see if it improves, it might be that 4% PFA is masking some epitopes, and so far I've never changed the PFA fixation conditions. I've tried the methanol before in PCFs, and as you say, the morphology is not very nice, I found that the nucleus loses its nice outline (DAPI) and appears to be leaking, which for imaging nuclear proteins is not great (the signal appears too diffuse). Thanks again! Best wishes!

normally reducing pfa concentration also reduces background and will leave you with stronger signals. you could check to use the triton together with the pfa fixation. this sometimes increases antibody accessibility. add 1/5 of your blocking reagent concentrations to your antibody solutions (we use PBS, 0,5% cold fish skin gelatine) and increase the salt concentration to 0.5M NaCl - the latter reduces non-specificity.

josef

Hello Catarina,

I've never worked in this field, but as your marker need to go through several membrane to reach its target, it might be worth to go with the smallest fluorochrome available for your target, in order to increase its chances to reach the nucleus.

You can find a table with the corresponding size of the different generally available fluorochrome at http://flowcyt.salk.edu/fluo.html.

As mentionned before, could you check that the fixation protocol does not alter the epitope ? Is there any "positive control" where your epitope would be found at the cellular surface ? You could then try to fix these cells with your procedure and make sure the epitope is still efficiently detected post-fixation...

Hope this can help.

Fred

Hi Catarina,

I also never worked in your field, but I improved my nuclear stainings with two different approaches:

First of all, you could do an antigen retrieval, which should be done in PFA fixed nuclei. I tried there several methods, but cooking the slides in citrate buffer for 10 min helped a lot.

To circumvent this procedure, you could also fix your cells in Aceton/Methanol 10 min on -20C, completely air dry and then permeabilize in 0.1% Triton X 100 for 10 min.

I also perform my stainings in the blocking buffer I'm using which includes 0.1% Triton for all following steps.

Good luck

Eva

What it was working for me I do the imunnostaning for isolated nuclei, because my antibody could rich nucleus. So maybe try this.

Hi Catarina,

I have worked with imaging of Nuclear proteins and I can imagine how frustrating it could be. Increasing T-X-100 is not the solution. What I have been doing is that I pre-warm PFA and T-X-100 at 37 degrees and do 20 min fixation at RT and T-X-100 permeabilization for 10 minutes. One more important step is that I incubate with 20mM Ammonium chloride which is an excellent quencher for paraformaldehyde that causes the background fluorescence especially in green channel.

Hope this helps.

Hi Catarina,

How confident are you that you have a good myc antibody for immunofluoresence? The few times I tried imaging myc-tagged proteins with immunofluoresence I had terrible results. Do you have any other markers that you can try labelling with? If other antibodies label well in the nucleus, I would suspect that it's your primary antibody that's the problem.

Good luck!

Hi Catarina,

I do a lot of IFA on nuclear proteins of T. brucei. I have the best experience with HA and v5 tags. I also have one line with 13 x myc tag. The signal is not so bright as with HA or v5 but still OK. I have been also using 3 or 4% PFA (in PBS, 1.25 mM NaOH and heated to 60 and vortexed occasionally until the PFA dissolves) for fixation (couple of minutes on ice in the eppendorf and 10min on the slide). Then I wash once with PBS. I do permeabilisation either with 0.2% Triton in PBS for 5 min at RT, OR with 100% methanol in the freezer (-20, the MeOH was pre-cooled) for 20 min. Both worked fine with me but I mostly use the 0.2% triton for permeabilisation. Then wash once in PBS and block for 1h RT with 20% FBS in 0.1% Triton in PBS. The antibody incubations also 1h RT in 20% FBS/0.1% Triton/1xPBS. I think that the presence of 0.1% triton in the antibody incubations might help the antibody to get inside. I do 3 x 5min washes with PBS after both the primary and the secondary antibody incubations and final wash with milliQ water after the 3 PBS washes after the secondary ab incubation to remove the salts. I use the prolong gold mounting solution with dapi, which is hard set and the signal is brighter than with the ordinary one. I hope this is helpful.

cheers

Ludek

Hi Catarina,

I agree with people, IF with nuclear proteins in Tryp can be frustating. My two cents: when I was working with IF, I usually tried different detergents and concentrations for permeabilization. For example, first try would be Triton-X 0.5%, NP40 1% and NP40 0.1%, 20 min at RT: usually NP40 rendered me better results.

Fixing was never a major issue in my hands: 2% PFA for 30-40 min on ice, 1% PFA o/n, 4% PFA for 20 mini, but always ice cold to get the fastest fixation.

Also, I agree with Chris about how good is your Ab: not all Ab are good for all techniques. It could be worth to check if with similar protocol you can detect other nuclear proteins.

Best wishes and good luck!

Hi Catarina,

I agree with many former posts, specifically alternative tags such as V5 which has worked marvelously as a nuclear epitope for me in the past. As a second, easier alternative you may consider using a red fluor secondary (Dylight or Alexa) as these tend to have better signal to noise that green fluors under higher power such as confocal scanning where less signal normally observed.

Lastly, relative to fixation - for my nuclear staining (which was not in Tryp cells) I use 10 minute ice-cold ethanol permeabilization in lieu of triton after 4% PFA fixation. In addition, if you are having background issues with PFA 1) use fresh (less oxidized aldehyde groups) and 2) you can try to quench free aldehyde with 50 mM glycine/lysine combo (or 100 mM glycine if that's all you have).

Best luck!

Hi Catarina,

If you haven't got your heart set on IF you could always try a subcellular fractionation method followed by immunoblotting- may also be a good technique to use in parallel to IF to validate findings.

Good luck!

Hi everyone,

Thank you all for your various replies! I'll take your advice and try several combinations.

Thanks again!

Best,

Cat

Hi Catarina,

Just after Triton treatment, you could also try to plunge your sample into liquid Nitrogen: Incubate in PBS 1X, 20% Glycerol for 30 min, then freeze in liquid nitrogen for approx. 30 sec and thaw. Repeat this step twice, i.e. three freezing/thaw cycles in total.

This is what I do for drosophila cells to help large DNA probe to penetrate nuclei. Good luck. Fred

Hi Catarina,

usually, diffusion in cells is limited by particle size and mesh size. That means, firstly, you should try to use the smallest possible molecule for staining, i.e. possibly antibody fragments instead of complete antibodies and small fluorochromes (avoid fluorescent proteins). Secondly, you should avoid too much of crosslinking by formaldehyde (or PFA). We preferred fixing in 1% formaldehyde plus 4,5% acetone to damage cell membranes (and improve fixing) - at least that worked well in human cells. We even did that without triton. Maybe that works in Trypanosomes as well.

Good luck !

@ Frederic Grosjean

The fluorochrome table you linked is great !

My favorite "all compartment" fixation is to use 4% pFA in PBS for 15 min, followed by -20 methanol for 10 min. The methanol both fixes and permeabilizes the cells, allowing better access of the antibody. Using just methanol alone flattens the cells and compresses the nuclei, so that is not good--but the combination works very well. Good luck!

I agree that fixation using formaldehyde should be avoided if possible to avoid the cross linking problems. Have you any access to x-ray diffraction equipment?

Although x-ray diffraction will not give direct imaging it should allow information to be gathered either from the protein in situ if the diffraction angle is known or from a solution. Resulting data can be reconstructed into an image.

The advice given before gives more information on pre treating cells that could be followed (hopefully avoiding formaldehyde if using diffraction techniques but not necessarily a problem). Use of cryogenic freezing is a good technique to use to image however things are done.