Hi Van Thai Than,

There are lot of posibilities for fixatives. Paraformaldehyde 4% (prepared in PBS buffer) for 10 minutes at RT  it will be a good start  point, you may adjust time and temperature then to your set up. Glutaldehyde 2.5% could be also. If you do not have those, also cold methanol (put it 10 minutes in the freezer -20°C before use) will work, but remember that Methanol fix structures and permeabilize membranes at the same time (PFA and GLA only permeabilize).

best regards,

Peter

Dear Dr. Peter Hans Döhmer,

Thank you for your interesting answer. I will try to optimize the experiment as you suggested.

Best regards,

Thai

Dear Dr. CI Falcon-Rodriguez,

Thank you for your supported protocol.

Best regards,

In my works, I fixed cells with 4% paraformaldehyde in PBS pH 7.4 (filter 0.45um) for 15 min at room temperature and wash the samples twice with ice cold PBS and follow the standards method.

You can use natural buffered formalin or 2-4% PFA in PBS. Fix cells for 15 min at RT.

I would suggest you to look at the primary anti-body catalogue for the compatible fixer. You can try 3-4% paraformaldehyde in PBS for 10-15 min at room temp. Also, you can try methanol as fixer for 30 min - 1hr in this you don't need permeabilization step. I thin 3-4% paraformaldehyde should work fine.  

Here is the protocol from my lab (attached document)

IMMUNOFLUORESCENCE ANALYSIS OF CULTURED CELLS: updated August 28, 2015

PROTOCOL – CITI LABORATORY

-      Materials needed: Cells, fixatives, antibodies (primary and secondary), curved EM tweezers, hooked needle, parafilm, plastic boxes, glass slides, glass coverslips, mounting medium, PBS (see also Recipes).

-      Using curved EM tweezers, place one sterile round glass coverslip (Fisher Microscope coverglass, cat n. 12-545-80, 12 mm circle) in each well of a 24-well plate. Sterilization is done by dipping the coverslip briefly in 100% ethanol and flaming it just before putting it into the well (alternatively, a petri dish of coverslips can be autoclaved, but this often results in coverslips sticking to each other). Comments: Use of round coverslips in 24-well plates is recommended for multiple stainings, multiple conditions etc. Do not waste 1 plate for 2 coverslips (better use square coverslips in this case (see below).  Do not squeeze coverslip too hard, take only 1 coverslip at a time. Coverslips can also be sterilized by autoclaving, but that can only be done for few coverslips, since they tend to get wet and stick to each other If >24 coverslips are needed (screening for monoclonals etc) one can also use multiwell glass slides, inoculated the night before with one drop of trypsinized cells (do not cover whole slides with medium unless there are many cells or they are left for a long time). Square coverslips can also be used. It all depends on the number of samples you want to examine (think before!).

-      plate cells on top of coverslips at appropriate density and grow until desired. Comments. You should plate enough cells so that at the time you plan to do the experiment there are enough but not too many. Sometimes cells need to be treated with drugs for a certain length of time, so take this into account. Also, different cells grow at different rates.

-      Remove medium by aspiration. Wash 1x-3x with 1xPBS (cold if methanol fixation follows). Use a 10 ml pipette, it allows better volume control than a 25 ml pipette. Use a blue tip at the end of the suction tube, it allows faster aspiration of liquid. From now on, be careful never to leave cells without any liquid on them for any significant length of time. When you aspirate, always leave a trace of liquid that covers the cells, to avoid drying them.

-      Fix cells following one of the procedures below. During fixation, prepare primary antibody solutions.

-      Place a sheet of parafilm on a perfectly flat surface (wax tablet, glass plate, flat bench, cover of 24-well plate), and just before fixation is over, place droplets of 30 ul primary antibody in neat rows, corresponding to positions on the plate, at enough distance from one another so that the coverslips are separated by  ~ 1 cm  (at least).  You can flatten the parafilm on the bench (if it is flat…) or on a support, such as for example a wax tablet. You should also label the parafilm or the surface under the parafilm to be sure which droplet/coverslip is which. It is very important that the parafilm be FLAT.

-      Remove coverslips from plate, and place upside down on droplets (cells on the side of the droplet). To lift the coverslips, use with one hand a needle with the very last part of the tip (0.1 mm) bent, so it forms a tiny hook, and with the other hand the curved EM tweezers, that you will use to quickly grab the coverslip once it’s lifted. You can also do it using tweezers alone (more difficult): press tips together, slide them gently UNDER coverslip and raise it slightly at an angle (10-30 degrees); release pressure on tips and gently slide the upper tip on top of coverslip, while maintaining the bottom tip under the coverslip. Re-apply pressure to clamp coverslip.  Be gentle with coverlsips, otherwise they break!! Take coverslip out of well and place on droplet, with cells obviously facing the liquid (down), and not the air.

-      Once all coverslips are placed on droplets, place them all in a humidified environment. This is done either by having the wax tablet+parafilm in a sealable plastic box that contains, on the side, a compressed, wet kimwipe, or by placing a box lid on top of the bench, and leaving a wet compressed kimwipe inside. Be careful not to disturb coverslips while doing this. Place the whole thing in a safe area, to avoid hitting the coverslips and mixing up the antibodies.

-      Incubate at RT for 60 min (or 30 min at 37, or 45 min at 30°C).

-      Transfer coverslips, cells UP, in a new 24-well plate (it can be reused for washes only), containing 1-2 ml per well of cold PBS. Incubate 5-10 min. If you are using square coverslips, use appropriate white ceramic rack for washes. If you are using large slides, use Coplin jars.

-      Repeat the washes 2x, either by transferring into new plate, or by aspirating PBS and adding fresh PBS. If you do that, be very careful and gentle adding PBS, since cells could be detached. Use a pipette-boy that does not have a too strong pressure.

-      During last wash, prepare sheet of parafilm with droplets of secondary, fluorescent antibody. Transfer coverslips on droplets, (cells DOWN) paying attention to putting the right coverslips on the right droplets. Incubate in a humidified environment for 30-40 min. Cover with aluminum foil, to avoid bleaching of the secondary antibody. Wash as above.

-      During last wash, label glass slides with date, cell type and antibodies used (or else, a number). It’s very convenient if you label the name of the antigens with the color in which they are stained (red for Cy3, green for Alexa488 or FITC, etc).

-      You can place 2-3 coverslips on each glass slide. Place 2 tiny droplets (2-3 ul Vectashield, typically) of mounting medium at equal distance from the edges of the slide, avoiding to leave air bubbles within the medium. It’s very important that you do not put TOO MUCH mounting medium, otherwise you will not be able to seal the coverslip, and coverlips will float, cells will detach or move , etc.

-      Using tweezers, remove coverslips from well, touch the edges gently with a kimwipe to remove excess liquid (do not touch cells!), and place coverslip (cells DOWN) on top of droplet. Leave 3-5 minutes to allow DAPI from mounting medium to penetrate evenly in all cells. Then apply a very very gentle pressure (with a yellow tip) to flatten the coverslip on the slide and to remove air bubbles trapped within the coverslip and the medium. Aspirate excess mounting medium from the sides, using air vacuum. Seal with nail polish, by applying a thin layer of nail polish between the edge of the coverslip and the slide. A good tip is to place first a tiny drop of polish at the 4 poles (N, S, E, W) of the coverslip, to fix it in place, then do the sides. Another mounting medium that can be used is Pro-Long AntiFade kit, that hardens after drying (so no need for nail polish). However use this only for very important or unique samples, since it is very expensive. It’s critical that a complete all-round layer of sufficient amounts of nail polish is placed around the coverslip, to seal it COMPLETELY from the outside, and BLOCK its movement completely.

-      Leave slides to dry the polish at RT for at least 3 hr, and then observe in the microscope. Never place under microscope slides where the nail polish is still wet: it will destroy the objective! Store slides in the cold. This is particularly important if the mounting medium is Vecta-shield (if you leave them at RT, the next day there will be nonspecific fluorescence on the sample).  If slides have been stored at 4°C, they need to be pre-warmed before going to the microscope, to avoid condensation of humidity.

NOTES ON ANTIBODIES

1)   PRIMARY ANTIBODIES

Before using an antibody, check that it works on cells of that species. Another good check is to see first if the antibody works by immunoblot, Secondly, if the antibody has a spec-sheet, check at what dilution it should ne used for immunofluorescence. Typically, the IF dilution is 10X less (eg 10X + concentrated) than the dilution used for immunoblotting.

Different antigens may require different fixation protocols. So, check on the spec-sheet if there is any indication on the preferred fixation. If this info is not available, check on the literature to see what fixation people use to see a certain antigen by immunofluorescence. Usually, we find that in most cases for TJ proteins MeOH fixation works very well. However, actin for example requires paraformaldehyde-Triton. Some sensitive antibodies (for example ZO-1 rabbit) require 1% PFA, and they will not work, or work less well, with 2-3% PFA.

The quality of the IF results depends primarily on the quality of the primary antibody.

2)   SECONDARY ANTIBODIES

We use Jackson secondary antibodies, that we store as a 1:1 glycerol suspension at –20°C. The final dilution is typically 1:100 or 1:200. If you think the signal is too strong or too weak, or there is too much background, you should do a test with serial dilution of secondary antibody, and use the highest dilution that gives maximum signal.

For confocal analysis, it is important that the fluorophores do not have much overlapping absorption/emission spectra. The best combination is

Alexa488 + Cy3 + Cy5

NB Cy5 is not visible under the eyepiece of our inverted Zeiss microscope, you can only see it with the laser line.

If you absolutely want to see the red signal in the microscope on your samples, you can also use FITC + TRITC, but you must make sure that the green does not excite the red (by turning off red lamp and looking in the red channel if there is signal) and if so adjust the filters and absorption wavelengths to abolish the bleed-through.

USE A POSITIVE CONTROL WITH A FIRST ANTIBODY THAT YOU KNOW WORKS.

FIXATION PROTOCOLS

1.    COLD METHANOL

Works well for most antigens, but not for actin.

Several hours before the experiments (better overnight) place a tightly sealed bottle with ~ 100 ml methanol in –80°C freezer.

Rinse cells with cold PBS(+Ca/Mg) 2x or 3x

Just before adding to cells, remove methanol from –80. Place 1 ml/well (24 well plate). NB If cells are on racks of coversips or on multiwell slides use a jar with methanol).

IT IS CRITICAL THAT THE METHANOL BE VERY COLD AND NOT HYDRATED.

Incubate coverslips at –20°C for 5-30 min (Geiger kidney primaries: 10min)

Wash 3-4x with PBS and leave in PBS for at least 10 min to rehydrate.

Change the MeOH stock at least once a month, to make sure it’s good (not hydrated). Do not leave the MeOH bottle open for any length of time besides what is strictly necessary to get out the volume required for fixation. Leaving the bottle open will lead to hydration, and worse performance.

[Variation : methanol-acetone protocol. Just after incubation in methanol incubate in acetone pre-cooled at -20°C for 10 min).

2)   TRITON-PARAFORMALDEHYDE 3%

Protocol n.1

Rinse cells 3x with PBS(+Ca/Mg).

Permabilize/fix cells with 3% paraformaldehyde and 0.2% (formerly 0.5%, now reduced) TritonX-100 in PBS at RT for 2-3 min.

Postfix in 3% PFA in PBS (pH 7.0-7.2). Incubate 20 min RT

Wash 3x with PBS.

If you are using a frozen stock of paraformaldehyde, check pH before using!

Protocol n.2.

Rinse cells 3x with PBS (or with 50 mM MES-morpholino ethan sulphonic- pH 6.0 diluted from stock 0.5 M stored at 4°C)

Permabilize cells with 0.1% TritonX-100 in PBS (or 0.5% in MES, freshly diluted from 10% stock)) at RT for 2-3 min.

Remove all Triton by aspiration (or wash 5-6x with PBS)

Immediately add 3% PFA in PBS (pH 7.0-7.2). Incubate 20 min RT

Wash 3-6x with PBS

Protocol n.3

Rinse cells 3x with PBS

Fix in 2% PFA in PBS, 30 min RT

Rinse with PBS

Permeabilize with 0.1% Triton X-100 in M Buffer (50 mM imidazole, 50 mM KCl, 0.5 mm MgCl2, 0.1 mM EDTA, 1 mM EGTA, 1 mM b-mercaptoethanol, pH 6.8), 4 min RT

Wash with M buffer

Rinse with PBS + 0.2% BSA. Antibodies in PBS+BSA 0.2%

3) METHANOL (or ETHANOL)

Method for MDCK cells, from K. Matter

Permeabilize for 2-3 min at 4°C with 0.2% Triton X-100 in 100 mM KCl, 3 mM MgCl2, 1 mM CaCl2, 200 mM sucrose, 10 mM hepes pH 7.1

Fix for 5 min in MeOH at –20°C or for 30 min with 95% ethanol on ice

(alternate directly in ethanol for 30 min on ice, followed by 1 min RT acetone)

4.    PARAFORMALDEHYDE-METHANOL (C. CHAPONNIER)

1% paraformaldehyde (in PBS?), heated to 37°C

add to cells and incubate at RT for 30 min

Rinse 2x with PBS at room temperature (5-10 min each)

Incubate with cold methanol (kept at -80) at -20°C for 3 min

Remove ½ of MeOH, add same amount of PBS.

Repeat step above, so that you transition gradually to PBS

Remove all and rinse 2x with PBS. Leave for 5 min to rehydrate.

Proceed with staining with antibodies.

Lower PFA concentratiuon is required for certain antibodies (eg ZO1 polyclonal)

5.    PARAFORMALDEHYDE-METHANOL (MICROTUBULES- SERGE)

N.B. cells must not be completely confluent otherwise the permeabilization does not work properly

 Wash quickly 2X in PBS +CaMg by aspirating the medium and adding the PBS immediately.

Add the paraformaldehyde 3.7% (preheated at 37°C) and incubate at RT for 1 hour.

Wash quickly 2X with PBS

Add cold MeOH (-20°C) for 5 minutes

Wash for rehydratation carefully (4 times) (PBS can be added even before MeOH is completely removed)

prepare the antibody (30 ul per slide)

Incubate 45 minutes at 30°C

3 rapid washes with PBS

prepare the second antibody (30 ul per slide)

Incubate 30 minutes at 37°C (do not incubate more than necessary to avoid background)

Make 3 rapid washes

Mount the slides. Remove the drop present on the slide to avoid diluting vectashield.

6.    (PLP) PARAFORMALDEHYDE 1% / LYSINE /PERIODATE

In general 1% PFA (rather than 4%) is much better for a large group of proteins (eg certain ZO-1 antibodies)  and fluorophores and is as simple to make.

1% PLP (paraformaldehyde-lysien-periodate) is an extremely good fixative for almost every single antibody as it combines the ample but not overdone fixing power of 1% PFA, with a background/fluorescence decreasing properties of lysine and periodate. 

-----

To make 100 mls PLP, dissolve 1 g PFA in 60 ml of 60C dH2O. Solubilize with a few drops of 1 N NaOH on low heat and stirring.

Weigh out solid .1M Phosphates: 1.014 g Na2HPO4 (dibasic) and .392g NaH2PO4 (monobasic) and add it to the 1% PFA with stirring.

Simple paper filter it, and add to it 1.37 g of lysine mono-HCL, predissolved in 10 ml H20.

Finally, add in .21g Na Periodate and QC to 100 ml.

(this is very laborious, but it is an excellent fixative for people who are doing careful IHC, DAB or fluoresence in tissue. Also, the lysine and periodate appear to do some kind of complexing reaction, so the fixative needs to be made fresh, at most 6 or 7 hours before use).

7.    GENTLE PERMEABILIZATION PROTOCOL WITH SAPONIN (GRUENBERG LAB)

Triton X-100 is too strong a detergent, it will explode internal membranes (endosomes, etc). A gentler protocol involves fixing cells with PFA, followed by incubation of primary  antibody containing saponin at a concentration between 0.01 and 0.05% (Sigma S7900., 25 g)

They use a 10X PBS stock that also contains 10% BSA and 0.1% saponin, and they dilute this stock 1:10 to obtain the antibody diluent.

RECIPES

1.    Paraformaldehyde

-      heat ~ 80 ml PBS to ~70°C and add 3 gr paraformaldehyde while stirring; mix until clear

-      add 100 ul 0.1M CaCl2 and 100 ul 0.1M MgCl2 (to give a final concentration  of 0.1 mM) with stirring while the solution is warm, to prevent precipitation

-      allow to cool; make up to 100 ml with PBS and adjust to pH 7.4

-      store at –20 in aliquots and use fresh each day. Check pH after thawing.

2.    Mowiol mounting medium (home-made)

Mowiol 4-88                    2.4 gr

Glycerol                           6 gr

dH2O                               6 ml

0.2 M Tris-HCl pH 8.5     12 ml

Place glycerol in 50 ml disposable conical centrifuge tube. Add Mowiol and stir thouroughly. Add dH2O and leave for 2 hr at RT. Add Tris and incubate at approximately 53°C until the Mowiol has dissolved (takes years); stir occasionally. Clarify by centrifugation at 4000-5000 rpm for 20 min and aliquot the supernatant into glass vials with screw caps, about 1 ml each. Store at –20°C, stable at this temperature for 12 monbths. Once defrosted, stable at RT for 1 month.

Elvanol: mounting medium:

(16.7% Mowiol, 33% glycerol, 0.1% phenylenediamine)

3.    Vectashield mounting. Use the Vectashield product. Use 5 ul for small round coverslips, 10 ul for large coverslips, one drop for sections on slide, then place coverslip on top, then seal the edges with nail polish. It’s easier to do with large coverslips.

4.    Airvol mounting medium for immunohistochemistry sections.  Dissolve 10 gr of Airvol (polyvinyl alcohol, Air Products, Inc., Allentown PA) in 40 ml 50 mM Tris, pH 8. Stir overnight at room temperature to dissolve. Add 20 ml glycerol. Aliquot into 10-ml syringes and store at -20°C. Thawed aliquots have a shelf life of 2 weeks at room temperature.

5.    Glycerol anti-fade mounting medium (90% glycerol with n-propyl gallate). Recipe for this is:

- Prepare a 10X PBS stock solution.

- Prepare a stock solution of 20%(w/v) n-propyl gallate (Sigma P3130) in dimethyl formamide or dimethyl sulfoxide. (Note: n-propyl gallate does not dissolve well in water-based solutions.)

Thoroughly mix 1 part of 10X PBS with 9 parts of glycerol (ACS grade 99-100% purity) and slowly add 0.1 part 20% n-propyl gallate dropwise with rapid stirring. 

6. DAPI. Make stock 5 micrograms per ml, dilute 1:1000 in secondary antibody.

TROUBLESHOOTING

-      Coverslip problems (breakage, stuck, double, etc).

Causes/Remedies

a)   Coverslips got too hot during flaming. After the coverslip (dipped in ethanol for sterilization) ignites, immediately wave your hand fast enough so that the flame burns all the ethanol quickly without overheating the coverslip. Handle the coverslip gently with tweezers. Never squeeze too hard, the coverslip will break.

b)  Coverslip placed in well while too hot. Allow flame to extinguish before you place coverslip in well. Otherwise it will melt the plastic and fuse to the glass.

c)   Two coverslips placed in the well. Pay attention when you take coverslips from stock. You should only get 1, not 2-3. Even if you have more than 1, you can still be able to retrieve the top (good) coverslip, and remove the 2nd, unwanted one, and proceed with the staining.

d)  Square coverslips overlap. When using square coverslips in a dish, they can move and go one on top of the other. As a consequence, coverlips will have only part covered with cells. Avoid that by separating coverslips with a sterile pipette and moving dish gently after adding medium.

-      No or few cells on the coverslip. The cells were detached during the procedure.

Causes/Remedies

e)   Cells dying or dead. Monolayer too confluent. Make sure cells are not too confluent on the day you plan to fix and stain them. If they have to be confluent, make sure they are fed the day before the fixation. Look at the coverslips before you fix. If there are no cells, it means cells have died and disappeared already (in this case, it’s unnecessary to stain…). This problem can also be caused by improper handling of the cells: cells become sick when not trypsinized at the right time, when they let go overconfluent, or are seeded at too low density, etc. Cells must be cultured properly and watched every day!

f)    Cells scratched off the coverslip, or on wrong side of coverslip.  Make sure you always know for certain which side cells are on. Do not scratch with tweezers. Hold the coverslips from the sides, do not touch centre of coverlsips with tweezers

g)   Cells flushed away by solutions/buffers. During all steps of fixation and washes, make sure you add solutions to coverlips not directly on top, and not too fast. Otherwise the solution mechanically tears away the cells.

h)  Cells require collagen coating. Some cells will simply not stick to glass alone, they require collagen of polylysine coating. Transwell filters are very good to get good polarization of cells if does not work.

-      Antibodies did not seem to work. No specific staining.

Causes/Remedies

i)     Wrong first antibodies were used or inappropriate dilution. Make sure that for each primary antibody you use you know exactly its characteristics: is it rabbit or mouse? What was it raised against? Has it been tested before on this cell line? Does it work on this cell line? What dilution should it be used at?How old is the batch I have been using? Has anybody left this antibody at room temperature, or spoiled it otherwise?

j)    Wrong secondary antibody . If using a primary rabbit antibody, use a secondary antibody against rabbit, not mouse!  Check the dilution. Always use the lowest possible concentration that gives the strongest signal. First time you use a new antibody, test 3-4 different dilutions. Always use a positive control when using the antibody the first time.

k)  Wrong fixation (see below).

3)  Non-specific staining, bad staining, strange staining

Causes/Remedies

l)     Improper fixation. Fix cells precisely according to the protocol. Do not let cells on coverlips dry at any stage during the procedure. This means being rapid, efficient and precise in removing most but not all the liquid covering the cells (leave just enough so that they are wet), and replacing with other buffer rapidly. Also, incubate the coverslips in humidified chambers. Make sure the methanol is cold and not hydrated, the paraformaldehyde is fresh and at correct concentration, and that all solutions contain all they are supposed to contain and nothing more or less. In addition, some antigens require a specific fixation protocol, otherwise they are not seen or the staining is very bad. For example, actin requires paraformaldehyde fixation. If you fix with methanol, staining will be horrible. So check before hand whether you can use any fixation or a specific fixation with that specific antigen, If in doubt, first use methanol fixation, then repeat with paraformaldehyde (or others). The batch and quality and concentration of Triton is very important. The pH of the paraformaldehyde solution is also very important.

m) Improper staining or washing. Use appropriate antibodies at appropriate dilutions. Incubate with first or secondary antibodies for the appropriate length of time. It’s better to shorten than to lengthen incubation times. So, keep incubation with primary and secondary antibodies between 20-60 minutes (depending on temperature) and duration of washes between 5-15 min.

4)   Good areas/bad areas in the slide.

Causes/Remedies

n)  Partial drying of coverslips. Make sure no part of the coverslip is dried during any step. Most often, this problem is caused when coverslips are partially dried during incubation with first or secondary antibody (coverslips not flat, air bubbles). Make sure there are no air bubbles between cells and antibody solution, and the coverslips are horizontal, flush with the support, in  a humidified atmosphere.

o)   Improper mounting . When mounting coverslip on slide using mounting medium, make sure there are no air bubbles in the droplet of mounting medium. Make sure that coverslips is gently pushed towards slide to remove excess mounting medium or air droplets.

Sandra Citi, MD PhD

University of Geneva

4, Boulevard d’Yvoy

1211-4 Geneva

Switzerland

[email protected]

Dear Prof. Sandra Citi,

Thank you very much for your detail protocol.

Sincerely,

It's a pleasure to share the experience of decades of mistakes (and successes) with young researchers...:)