Hi,

What are you using as secondary antibodies?

Anyway, you may try to label tissue wit Black soudan prior to proceed with the immuno.

Best,

Etienne.

get a more specific primary (monoclonal) antibody and consider increasing your BSA concentration to decrease non specific binding of your primary. Also hope your secondrary is not raised in mouse.

To block the tissue, I usually use serum from the species the secondary antibody come from, at 10%. If several species are involved, I mix several serum, but I usually use Donkey serum most of time. I agree that 2% BSA may not be sufficient. If the host species of your primary antiboy is the mouse too, you may use a M.O.M kit or an unlabeled Donkey anti-mouse IgG as you already said.

As Tumi Chabaesele said, try to increase the BSA concentration. In order to check non-specific binding of 2nd antibody I suggest a negative control (only aply 2nd antibody without 1st).

I concur with Etienne. I use the serum from the species in which the secondary body was generated. Check the concentration of the secondary antibody. Both, excess concentration and incubation time can increase non-specific staining.

I don't typically use IgG's to block with. In fact I typically use BSA (IgG free) at 5-10% overnight in the cold, or fish oil or low IgG skim milk. You say that you get a lot of background non-specific staining in the mucins, it may be possible that your antibody is trapped and not bound per se. Have you tried to increase washing or changing washing solutions? Also, Mr Krause suggested doing some background controls using only 2ndary antibody. Have you looked at that. Can you decrease your primary and/or secondary concentrations and still maintain a reasonable signal?

I have not direct experience with such a pathological tissue (may the mucus show strong auto-fluorescence? Try to check in negative control tissues without Ab), but I always found a solution to my lack of specificity trhough the most commons histological "tricks".......

Non-specific signal to other cells/tissue regions is a common issue with Immunofluorescent labelling (also known as background staining). Causes are various. But there are a number of well-documented precautions that can be taken during sample preparation and protocol execution, to ensure that non-specific binding is minimized.

First at all, I generally use in my blocking, a 5% BSA concentration (2% can be too weak to cover all non-specific sites)

Some general tricks:

1. Whe possible, employ direct immunofluorescence. Indirect labeling can boost overall signal intensity, but the use of secondary antibodies can result in extensive non-specific binding, compromising signal to background ratio - In case of direct IF, dilute the fluorochrome-conjugated antibody further. If it is too concentrated, background may increase due to an increase in non-specific interactions - In case of indirect IF, use cross-adsorbed secondary antibodies to remove unwanted cross-reactivities between labeled antibodies and any other immunoglobulins inherent or added to the experimental system.

2. Be sure to include a blocking step to reduce background by blocking non-specific interactions between the primary antibody and the cell surface or intracellular structures. I prolong the blocking step up to 3h at RT, when necessary (staining of little molecules, or few concentrated)

3. If possible, use F(ab')2 fragments where background may be due to binding of the whole molecules of primary or secondary antibodies to Fc-receptors. F(ab')2 fragments of most secondary antibodies are readily available.

4. Incubate fresh tissues or cells with normal serum in the presence of sodium azide prior to addition of the primary antibody.

5. For fixed tissue, remove free aldehyde groups with 0.1% sodium borohydride in PBS for 30 mins prior to staining.

6.In case of auto-fluorescent components, I also adopted the antigen retrieval (AR, you can easily find acid- or basic- AR protocols online) to swhitch off the non-specific signal, deciding for the most appropriate one (acid/basic) simply by experience.

7. The cause of non-specific staining can be due to too long incubation time (check under microscope every 1/2h!), or too high concentration of I Ab (try eventually a dilution curve, such as 1:1000, 1:500, 1:250, 1:100), or too high temperature (i.e., try to move from RT to 4°C: cold temp. slows down the reactions, increasing interactions specificity)

8. Use secondary antibody that will interact with primary antibody, antibodies that are raised against the species in which the primary was raised. For example, if primary antibodies are raised from rabbits, use anti-rabbit secondary antibodies.

Treat sample with e.g. Mouse-On-Mouse blocking reagent prior to the primary antibody incubation.

9. Avoid drying out of the samples, reducing the covering surface by using specific pens (PAP Pen) to draw a hydrophobic circle around slide-mounted tissue

10. For fixed tissues: try to reduce fixation times as much as possible

Helpful links /references:

www.protocol-online.org

www.ihcworld.com

Let me know.

Best,

Eva

Thank you all for fast feedback. All our primary Abs are not in mice, but in rat, hamster, rabbit or goat. We get non-specific staining irrespectively on the staining procedure that we use (primary labeled Ab, primary - secondary Ab, or primary Ab labeled with biotin and streptavidin). Avidin/biotin blocking was done with Endogenous Biotin/avidin blocking kit according to the recommendations of the manufacturer, so we do not think that is problematic.

To summarize suggestions so far:

1. We should increase BSA concentration

2. Add more blocking IgGs from different species (we use goat anti-rabbit secondary Ab, so we do not add rabbit IgGs for blocking)

3. Try blocking overnight at +4C (at the moment it is 1hr on room temperature)

4. Improve washing (prolong washing time and number of washes)

Hopefully this will help.

To me, this sounds like an autofluorescence problem. I have attached a protocol here that I obtained from Colleen Manning (in Jim Trimmer's lab at UC Davis). They employ Sudan Black (as one other person suggested already) to eliminate autofluorescence in immunofluorescent labeling. As requested in the attachment, please be sure to cite the protocol as Colleen asks.

To Etienne,

Thank you for the suggestion to use Soudan black. Do you have a protocol or a reference that we might try? Does it interfere with endogenous GFP?

sorry- hit the wrong button. Here's the attachment for the Sudan Black protocol from Colleen Manning described in my previous answer.

How thick are the sections you are using? A method I commonly use in tissue sections which I noticed was not mentioned here was in addition to increasing the concentration of your protein in your blocking buffer and increasing the blocking time to overnight is to include a small amount (0.1-0.3%) of detergent such as trition-x in all your regents. This helps that blocking proteins and also your primary and secondary antibodies to permeate your tissue sections.

Autofluorescence in lung tissue is a notorious problem. I'd be curious to know if the sudan black trick works. I seem to recall from seeing other people struggle with this, that it's worse in the red channel? Can you swap what colors you're using for which probe, and minimize the issue that way?

In general, I agree with the people who said to use serum to block. The protocols on my website are for IF on cell culture, but can be modified to use with tissue.

Raising the incubation temperature to 37 degrees increases stringency and reduces background for all antibodies.

I also agree with people's suggestion to try different secondaries. They're not all created equal. Sometimes it's a bit of a trade-off between signal intensity and background crap - the cleanest antibodies are sometimes dimmer fluorophores, but you can adjust the ratios (1:100-1:1000 depending on the fluorophore and your imaging system).

I didn't see whether anyone suggested pre-absorbing the secondaries yourself - if you can afford enough material to do this, it can help a lot. My website is http://samzeitlin.com and the protocol is there under "frequently requested protocols". feel free to contact me if I can be of any further help.

IF is just chemistry - if you understand the principles of each step, you can modify to optimize for your applications. best of luck!

Something I noticed was that the slower the blocking stage, the better the outcome. For example, blocking overnight at 4 degrees was better than a few hours at room temperature. Might help. Good luck

The problem sometimes is that the mucus trap antibodies. That is something you can not avoid because you are working with an asthma model. I noticed you say that you are using goat serum to block your slides. Goat serum is very sticky and is not recommended for blocking non specific binding. Normally, I am trying to buy secondary antibodies from Jackson Immunoreseach made in Donkey. They have good antibodies for multiple labeling that are adsorbed with serum from different species. Thus, I am alway blocking with Donkey serum. They also sold good quality donkey serum. It has to be clear, if not that can give you problems during the stain. If it is possible you can try to use Fab antibodies or use Fcblock (anti CD16/CD32) in your blocking step. In paraffin sections, it is good to add 0.1% Tween-20 and 0.1% Triton-X-100 to the PBS for dissolving the serum and the antibodies. I block with the serum only for 30 minutes, and I dissolve the primary and secondary antibodies in PBS with 0.1% Tween 20 and triton, without serum. You can also titrate the antibodies to see if you can reduce the non-specific stain.

For the collection, it is always good to bleed the mice to prevent accumulation of auto-fluorescent red blood cells in the lung. Perfused the lungs through the trachea with 10% neutral buffer formalin and do not over fix the tissue. 48-72 hours is enough.

What molecule are you trying to detect in the lungs? Sometimes is good to check in biocompare for antibodies that are good for IHC. Look for those that work in formalin-fixed, paraffin embedded tissues, and look for convincing pictures.

Javier

I have to agree with the other posts. It could be an autofluorescene issue. You can resolve this by including the following controls: an unlabeled control i.e., no antibody (just the buffer/ solvent it is diluted in) and a separate control with your primary antibody, that has been incubated with a respective antigen to block it's binding sites. Do your normal IHC protocol with these controls and observe if autofluorescence or non - specific binding is the issue. If autofluorescene is the problem, try and figure out how to remove it by playing with the gain function on a confocal, or even better, what wavelength is causing the fluorescence. Then you can just get a very specific label that fluoresces outside of the autofluorescene range. I hope this helps! You can also try another blocking agent, such as serum instead of BSA. Be careful of using too much, it could over block and prevent any signal from appearing, except for the autofluorescene.

Dear Berislav Bošnjak,

I just used Sudan Black one time. The whole fluorescence was decreased.

Here's what I did :

Unmasking (Citrate buffer 0.01M 90°C 30 min)

Rince 3 times 10 min

Sudan Black 1% diluted in EtOh 70% , 30 min 37°C

Rince 3 times 10 min

Quenching (Glycin 1mg/mL) 1h Room Temperature

Rince 3 times 10 min

Blocking BSA 4% + serum 10% 2 hours Room temperature. (depending of the thickness of your slices you can do it Over Night at 4°C) + Triton (if needed)

Primary antibody : ON 4°C with serum 1% and triton 0,1%

Rince 3 times 10 min

Secondary antibody (Alexa 1/500 2h Room temperature)

Dapi

Rince 3 times 10 min. (Over Night 4°C is better again)

I didn't read the protocol of Douglas Vetter... Hope it's quite the same! ;)

Find also in attached piece an other article on Sudan Black used for fluorescence microscopy.

Please, tell me if your results are better,

Best,

Etienne.

My suggestions:

1- Use signal enhancer: (EX: Image-iT® FX signal enhancer). It is a liquid that is applied directly to slides or coverslips containing fixed and permeabilized cell or tissue samples prior to staining with fluorescent probes. When Image-iT® FX signal enhancer is used, nonspecific fluorescence (background) commonly seen with the application of fluorescent conjugates of streptavidin, goat anti-mouse, or goat anti-rabbit IgG is largely eliminated.

2- Incubate with the primary antibody 3 hours at 37 C instead of RT. then overnight in 4 degrees.

3- Incubate with the secondary antibody 30 min to 1 hour at 37 C instead of RT.

I always do that and have less background and better images.

One thing that hasn't been mentioned here is to do your imaging in a space that is as dark as possible. A very common issue I see in fluorescent microscopy is the assumption that because one can distinguish the hot signal from the background, they are getting a good picture --- this is while allowing ambient light from around the scope to penetrate the sample. Ambient light has UV light in it and will stimulate autofluorescence. I think that with very dark imaging you get two benefits, the darkness helps the hot signal appear more, and the lack of background autofluorescence from ambient light allows the natural contrast to enhance.

Cheers

Thank you all for your suggestions. We are sure that it is non-specific staining of the mucous in the bronchial epithelium, as all non-stained controls or fluorescence-minus one controls are fine (6 um thick cuts of frozen lungs). For the moment we will try the new protocol with more extensive blocking (using 10% donkey serum) and more extensive washing (addition of Triton X-100 to PBS and prolonged washing time). We will see how this work, hopefully this will do the trick. For the moment we will not try out sudan black staining, unless the autofluorescence will be a problem.

Various methods we use. Incubate in 1% sodium azide to Quechua aldehydes, use the antigen retrieval method 0.1 M Na Citrate, pH 4 at 90 deg C (double good for antigen retrieval and in situ antibody elation/in activation, use preadsorbed second ( Jacson Immunoresearch), maximally diluted second, preferably fab fragment.

In the case I had a non-specific staining of primary antibodies in tissues sometimes was successful to re-use antibodies next time without changing the staining protocol (antibodies preadsobed with the same biological material). My protocols include what you have tried/planing to apply: 10% serum in blocking solution (before antibodies), Triton X-100 in all steps, long washes. Usually I use 0.25% BSA with both antibodies.

If, you want to stan mouse tissure, Do not use mouse IgG as blocking reagent. Thanks

You can treat the non-specific antibody with cold acetone.

Baptiste, can we use Zenon labeling method for thicker sections like 30 microns? 

Hi Berislav, I am currently having issues with autofluorescence in trachea tissue and think this is also due to the mucus. Did you manage to resolve your issue to get rid of the non-specific signals?

Cheers, Joanna

Dear Joanna,

Unfortunately not completely. We also got a lot of non-specific signals from mucus in major airways, especially with two-step staining procedures (e.g. Ab biotin - labeled streptavidin). We tried extensive blocking and signal enhancer, but they did not help and for certain markers (such as chemokine receptors) I had to give up. However, we now have simple staining procedure with directly labeled Abs without extensive blocking which works relatively well for well expressed markers such as CD4 (directly labeled Abs). Now when this works, I think that for all future applications I'll follow the advice from Baptiste and directly label the primary Abs for other markers. I hope this helps. 

Thanks Berislav!

Dear Berislav

I get a strong non-specific signal in endometrium glands filled of  mucus in mmunofluorescent microscopy . I would appreciate any suggestions on how you had got rid of these non-specific signals in your lungs.

Thanks!

i would use unrelated species IgG to block mouse tissues

for example a mixture of 5% horse serum, 5% goat serum, 5% FBS

also it depends on what secondary antibodies one uses based on that prepare your blocking reagent to avoid background from secondary antibodies.