I recommend you to fix the cells with 4% PFA: 

1. Aspirate off media from cells and wash with 500 µl 1xPB for 5min

2. Aspirate off PB and add 4% PFA to fix cells for 30 minutes at RT

4. Aspirate off PFA4% and wash 3x with 500 µl 1x PB.

5. You can continue with your ICC protocol or store the cells with PB 0.1M containing O,O5% azide in your 24 well plate. 

I recommend you to fix the cells with 4% PFA: 

1. Aspirate off media from cells and wash with 500 µl 1xPB for 5min

2. Aspirate off PB and add 4% PFA to fix cells for 30 minutes at RT

4. Aspirate off PFA4% and wash 3x with 500 µl 1x PB.

5. You can continue with your ICC protocol or store the cells with PB 0.1M containing O,O5% azide in your 24 well plate. 

Dear Emiri,

Easiest way to do this is to replace the culture medium for the formalin solution. 10% sound quite concentrated to me, I'd prefer 2%, though I don't know your specific settings. If your colleagues tell you 10% is fine, just do so.

So, keep the glass slides in the wells if possible.

·         Do you really need to take them out, you can use a needle tit a bent point and a pair of tweezers/forceps. The point of the needle (18 or 20 gauge) can best be bended by hitting a hard surface with the top. A small bend is enough. Catch the cover glass with the needle and take it out with a pair of tweezers.

Take away the culture medium

Pour in the formalin solution. 500 μl per well must be enough. Have the solution at room temperature. If the cellular process you want to study if vulnarible and heat/cold sensitive, do as follows. Take off some medium until 250 ul remains in the well, keep the plate 37°C. Pour 250 ul pre-warmed (37°C) 4% formalin solution in the well, yielding 2% FA end concentration. Do not wash the cells prior to the fixation. Washing before the fixation may start cellular processes that interfere with your goals.

Keep this for a couple of weeks at 4°C but do not freeze it.

If you want to stain the cover slips. Better to take them out than to let them in the wells. The reagent to stain with can bet dropped on a sheet of parafilm . Place the cover slip on the drop, cells faced down.

Dear Emiri:

You can fix your cells on a coverslip as follow:

You can make your own fixation buffer or you can buy(Parafolmadehyde solution,4% in PBS(MFCD00133991)

1. Fixation buffer preparation.

- 1ml 37% formaldehyde solution

-1ml 10X PBS

-8ml water.

Fixation of cells

- Aspirate  growth medium,and wash with 1ml of 1XPBS (cell culture grade)

- Add 1ml of fixation buffer

- Incubate at room temperature for 10minutes.

- Wash twice with 1ml of 1X PBS

- Add 2ml off 70% ethanol. Cells can be stored at 4C up to several weeks.

I hope this  will help you

Rachel Lokanga

Always prepare the 4% paraformaldehyde in PBS the same day you intend to use it. Warm but do not boil to dissolve the paraformaldehyde. (Do not breathe the vapors). When it is dissolved and cooled to room temperature, correct the pH to 7.4. This method results in the fewest artifacts.

Dear Rachel and Marian,

Can any of you give a reason why you wash the cells with PBS prior to the fixation? Most likely that is just because it is in the protocol. Please reconsider, lots of things can change during the washing step. What happens if you wash prior to fixation:

In the first place you wash away medium… nothing that is in the medium will mean harm to the cells and to the biochemistry that follows the fixation.

Many processes within the cells will change during the washing. For some this is not a big deal, though you never know what changes. Especially Marians 5 minutes washing is extraordinary. Many signalling pathways change really quick.

My advice is to keep as close as possible to the starting situation of the cells. You may want to remove some of the medium prior to fixation. At least leave a certain amount to cover the cells and know what the total volume of medium is. Add the same volume of buffered formalin at a double concentration (if the end-concentration should be 4%, start with 8% formalin).

“But the proteins in the medium will also be fixed”, who cares. They don't bite.

Of course you can think of situations where you absolutely will have to remove medium; e.g. when incubating cultured cells with Ab than need to be tracked in later stadia.

Good luck,

Duco

Over 20 year experience with culturing and immune staining.

[Apologize for lengthiness]

@Duco and Emiri (and  - perhaps - others): please keep in mind the following:

10% Formalin (R,TM of MERCK; maximal stable percentage of 'formalin solution' -commercially available - to be 37%-38% hydrous solution of the gas 'formaldehyde' in water)  

= 4 % hydrous formaldehyde solution (= e. g. using para-formaldehyde powder, you easily can produce a 1, 2, or 4% hydrous PFA-solution by yourself, always freshly used -  without any other stabilizer like in commercially available solutions - and achieved by warming up to max. 70°C and adding DROP by DROP 1 NaOH under constant swirling, until the solution becomes CLEAR).

[cf: https://en.wikipedia.org/wiki/Formaldehyde  or also:  https://pubchem.ncbi.nlm.nih.gov/compound/formaldehyde     [and far more citations and literature references) ]

Because (as this is also my experience here in RG) many young researchers don't know about that paradigma FA - Formalin(R),TM 

[=CH2O, (H-CHO) or H2CO-gas in water {theoretically} is soluble only until 40%, storable solutions therefore only 37-38 a% at the maximum! The latter only with stabilizer, usually +10%Methanol added and Ca-carbonate -in former days dolomite powder- to buffer / adsorb constantly 'produced'/formed/generated formic acid]

I recommend -  if you mention PERCENTAGES - always to define what you mean:  'Formalin' or, which chemically would be more correct 'Formaldehyde' since '4% / 8% formalin' is less than  '2% / 4% formaldehyde'....etc.

IMHO, Duco has given a practical hint on how to proceed (just to remove a certain amount of a known whole amount/volume of medium fluid in a well and replacing that withdrawn volume with the same volume of -now- double concentrated fixative (option: fixative formaldehyde[double concentration] in the same concentration / molarity of medium fluid or buffer used in your culture system. IMHO, 2% FA (and sometimes only 1%) in suited buffer solution for 5-10 min at room temperature [provided gentle shaking of plates] might be sufficient for fixing a simple cell culture...

But be prepared: such a short fixation with FA only does NOT STOP ANY/all metabolic reactivity in the cells (if cells then only are stored in a buffer/washing solution, some enzymes still function already afterwards, as we know this from ultrastructural studies)  for special tasks one could think also about another aldehyde (which unfortunately is more complicated to use due to its hazardous nature: acrolein...faster than FA).

Concerning washings: I agree with Duco that one should omit 'thorough' washing (or e. g. for '5 minutes') of cultured cells prior to fixation (possibility of artifactual metabolic changes...) and would like to emphasize the (in RG often read) paradigm which is often interpreted differently by the groups of "cell biologists/molecular biologists" vs. "morphologists/histologists":  Do not confuse: 

'PB' = phosphate buffer   vs. 'PBS' = 'phosphate buffered saline'.

Numerous Q&A can be found on this item in RG-archive (one has to search for but that can be accomplished really easily!). For space reasons/not to overstretch patience of readers and keeping lengthiness acceptable I won't go into detail here, but only would like to state (because this might be of weightiness:

1x PBS = concentration = usually "0.01M" (per se "hypo-osmolar" to cells)

10xPBS= concentration = usually "0.1M"  (per se ALMOST, BUT NOT really iso-"osmolar" to cells).

Which PBS-composition usually is applied by cell culture researchers usually is known by them or get the knowledge via existing and / or handed down SOP'S of institutions/labs. But there are several PBS-compositions (with different salts in different concentrations / amounts / volume) out in the wild, which might exert unexpected cellular reactions.

I would concur with Wolfgang and others. The different nomenclatures can be confusing: Formaldehyde; paraformaldehyde;  "formalin" ; NBF ("Neutral-Buffered Formalin"). 

Always best to assume two things that seem like they are the same thing, aren't. 

For my lab, we have always used 10% NBF as our fixative of choice. We use it for cell lines grown on coverslips or in chamber slides, cell lines that you scrape off, spin down and suspend in low-melting point agarose (a great technique for eventual incorporation into a tissue microarray!), and tissue samples. 

Why? I am a surgeon and its what I am used to in the OR with my surgical samples. Plus it is ready made, easy to dispose of with formalex (binds waste NBF and renders it non-toxic), and my fixation results roughly mirror what I will find in my human tissue samples when I bridge from cell-line or animal work to look back at a human cohort.

drain media (don't rinse). 15-30 minutes in 10% NBF (off-the shelf) formulation, then rinse in PBS and good to go. Don't store in NBF, you will over-fix the antigens if you do IHC.

Also, "10% NBF" I believe, contains 4% paraformaldehyde as the main working ingredient that does the fixing. Hope this helps not confuses.

Paul Weinberger

In my experience, you should better use freshly prepared 4% paraformaldehyde (warm 12,5 ml of distilled water until it is hot to touch, but not boiling, add 1g of  powder paraformaldehyde and a drop of 1N NaOH, stir utill the solution is clear. When it cools down, dilute 1:1 with 2xPBS. After fixing the cells wash them with PBS, then incubate in PBS with Triton X-100, and then with PBS +BSA.  Iwould perform ICH immediately after fixation and would not let the cells dry to avoid non-specific signal.