Phalloidin staining only at the edge of the coverslip

I have a strange observation and wonder if this forum can help to shortcut my troubleshooting efforts.
I stained cells grown on a coverslip with fluorescent phalloidin (DY-647), but for some reason only the outer edge of the coverslip shows a staining (see image below). There are cells all over the coverslip (see DAPI in the second image).
Here´s the protocol in brief:

  • fix with 4% PFA for 20 min
  • wash 3x with PBS
  • block with 5% NDS in PBSTx 0.05% (was a mistake, I intended 0.1%). approx. 30 min. [I know blocking is not necessary for phalloidin, but because others claimed the phalloidin not to work I wanted to include all steps]
  • Incubate with phalloidin in PBSTx 0.1% for 30 min
  • wash 3x with PBSTx 0.1 %
  • mount with ProlongDiamond. (the aliquot was probably expired - can this really go bad?)

Here is an overview over the coverslip. Phalloidin-actin staining only at the edge:

Here one can see that there are cells all over the coveslip (DAPI nuclei):
→ will send this image in a second post. As a new user I am only allowed to emed 1 image per post.

Does anyone have an idea what went wrong with the phalloidin?


Here is the image showing that there are nuclei in the middle part:


Weird! Did it only happen to the one coverslip? If so, I’d blame it on the lab gnomes.

Yes, weird indeed. Because people in lab have declared our “old” phalloidin as not working anymore we ordered a new one and for the first test of the new aliquot I used the new phalloidin in 2 different concentrations, the old aliquot, and a positive control from a neighboring lab:
All 4 coverslips show the same pattern.

So, either we have 4 lab gnomes working in synchrony, or it is something different.

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Hi Martin,
Ever tried a different fixative? In my hands phalloidin did not work several times with pfa fixation. Checkout adding Triton to the pfa solution, reduce the pfa concentration or best you check out using ice-cold MetOH. If all this doesn’t work order a new batch.

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Have you tried to image the center of the coverslip with my higher illumination and gain/longer exposure time to see if there is any staining there at all? My guess is that the outer edge of the coverslips have been allowed to dry during the staining. This often leads to a much brighter but less specific staining.

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This seems like perhaps a technical/coverslip processing thing. I have stained thousands of slips of PFA fixed cells with phalloidin. I understand that methanol risks diminished staining, as it changes the structural pitch of the actin filaments such that the phalloidin doesn’t bind as well. (This is not to say that phalloidin won’t work on methanol fixed cells, but I’ve heard it can be problematic). It does look like there is a gradient of intensity of the phalloidin as you move toward the center…Are you inverting the slip onto a drop of phalloidin, submerging it, etc., when you perform the stain? Your cells look pretty dense, are they piled up on each other? I always used 0.1 or 0.2% triton X-100. I never included detergent with the phalloidin, but this should be ok. Somewhat unrelated - is your DAPI in the ProlongDiamond?

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Thanks to everyone for the suggestions so far. To answer some of the questions raised:

  • DAPI is in the ProlongDiamond
  • I have the coverslips (18 mm diameter) cell side up on a piece of parafilm and fix and incubate with a volume of approx 100 µl. Washing steps with 2 ml volume
  • Possibilities that will be checked in the next testing round this week:

sorry … hit send inadvertantly.
Here is what I will evaluate:

  • fresh vs. old PFA (I was lazy and took a thawed aliquot I found in the fridge last time)
  • fresh vs. old Prolong (the one I had last time was kind of jelly-like)

I will keep you updated on the results.

yes, I had a close look at the center with higher magnifications. Pure black. No signal at all.

As mentioned in my post above, I do the incubation on the coverslip, cell side up, so the possibility that the border is not completely covered and might have dried out at some point is in therory there. But I stained a lot of coverslips in this way before … never had this issue. Admittedly, this was the first staining since a long time, but the technique feels to my like riding a bicycle - I don´t think I will unlearn this ever again :wink:

Hi everyone,
still very strange. I did some tests as written in the post above … all look essentially identical. Actin is only stained at the outer border of the coverslip.
This time I included a tubulin-antibody staining. Tubulin staining looks great on the whole coverslip.
Phalloidin647 was incubated together with the secondary antibody.

This phenomenon shows some consistency now, but it´s nothing I have ever seen in the last 10 years.
Any further suggestion is appreciated.

I have never heard of an edge effect on a coverslip. One surprising thing is that at the bottom of the image strip, one sees that some singular cells are stained further towards the center. Could it be that you have 2 cell populations? Maybe the cells at the edge differentiate? Can you use lifeact instead of phalloidin to check?

Med vänlig hälsning / Best regards



Sylvie Le Guyader, PhD

Live Cell Imaging Facility Manager

Karolinska Institutet- Bionut Dpt

Blickagången 16,

Room 7362 (lab )/7840 (office)

14157 Huddinge, Sweden

mobile: +46 (0) 73 733 5008


Of course I can not rule out anything, but last time I had IMCD cells and this time it´s HeLa. I don´t think they differentiate much. But who knows? The cells were seeded on the coverslip in a 12-well plate the day before I stained them.
I should probably have used an actin antibody instead of tubulin. Will do this next time.

Concerning lifeact - do you use this as a dye like phalloidin? I only know it as a lifeact.GFP plasmid. Maybe you are right and it is worth transfecting the cells next time.


Have you changed the mounting media? That would actually be my first guess, with something at the edge changing the chemistry (either the air or whatever you seal with).

100% glycerol is a great tool for this kind of troubleshooting. No bleach reduction, but also nothing that might make the fluors go weird. Side note, this is also a great option when trying to run down the source of autofluorescence.


I did not change the type of mounting medium, but I did use a fresh batch of ProlongDiamond + DAPI – no difference to the old batch.
I will include a glycerol mounted control in the next round. Thanks for the advice.

Any update? I hope no news is good news! I’m super curious since this kind of thing can come up in my core and it’s nice to be able to pinpoint areas for troubleshooting.

Unfortunately the problem is not yet solved. I promised myself to give it one more try before giving up. Did not take any pictures today but had a look at the microscope - depending on frustration level I will maybe take some images tomorrow. Probably not. I´m hoping for next week.

Today´s results in brief:

  • glycerol mounting (1):
    actin staining with an antibody (alexa488 channel) is nicely seen over the complete coverslip. Phallo647 fails. Almost nothing EXCEPT for some very nice cells in a very different z-plane (where there is no green to see). Can the glycerol RI induce such a strong abberation in z-direction? I was using a Zeiss Plan-Apochromat 10x/0,45.
  • glycerol mounting (2):
    cells transfected with Lifeact.GFP - they were glowing like crazy before fixing them. After mounting I see nothing. Also no Phallo647. Really no idea what went wrong here.
  • lifeact.GFP transfection (and ProlongDiamond mounting): very nice signal on the whole coverslip
    % when fixed with 100µl PFA top of the coveslip (18 mm diameter), there is the infamous Phallo647 ring at the edge of the coverslip.
    % when fixed with 500µl PFA in the 12well (i.e. coverslip completely immersed) I see no Phalloidin signal. Very strange. These are HeLa cells. They are not known for lacking the actin cytoskeleton, aren´t they?

Plans for the next round (more suggestions welcome):

  • so maybe it is really a drying effect that affect especially Phalloidin and not antibody staining and not GFP. Therefore: let one coverslip dry out on purpose. Any ideas at which step would be best? After fixing? After primary antibody? Before mouting? Currently I believe before mounting would be best.
  • maybe our HeLa cells grow really weird and have tons of F-actin when close to the edge of the coverslip and none when growing in the center? Therefore: break a coverslip before starting with the fixation. When half the coverslip has also an egde-staining, it can´t be the cells growing in the center are that much different (which I doubt anyways, the Lifeact.GFP looks very much the same, no matter where the cells grow).

Oh my gosh, this must be beyond frustrating. I’m pretty stumped myself. I cannot think of a reason the cells would survive with no f-actin. But…maybe something in your solutions is causing the actin to depolymerize before it’s fixed? Have you tried the age old replace-all-solutions-from-start-to-finish?

I never had troubles following the Ted Salmon lab IF protocol. If you aren’t working with microtubules you can eliminate the glutaraldehyde in the fix and skip the sodium borohydride quenching step. I can’t remember if the Hepes is for the MTs or the actin (it’s been many years)… Good luck! We’re rooting for you!

Problem solved!
It´s the 4%PFA. I did use a fresh aliquot in one of the former tests, but is seems all our current aliquots are not working. Might be a temporal coincidence with a change of the person in charge preparing the PFA.

What is weird though, unless you try to stain f-actin either with phalloidin or sprichrome´s SPY-probe (this was included in today´s test. Same behaviour as phalloidin) - you would not realize that the PFA is not OK. Antibody stainings (e.g. tubulin, actin) and transfected Lifeact-GFP looked perfectly fine.

Today I included 4% formaldehyde in the test: perfect staining on the complete coverslip. See images below.

Because a dried coverslip was also a posibiIity suggested above, I also included this: letting the coverslip air dry after the final washing step for approx. 10 min. Here I see no difference compared to my normal mounting method where I take of the liquid as good as possible but do not wait before mounting.

These are HeLa cells transfected with Lifeact.GFP. GFP in green, Phalloindin647 in magenta.
4% PFA:

4% formaldehyde:

I am still interested if anyone has an explanation what could possibly be wrong with the PFA.



I forgot to post one off the most puzzling images:

In this case the cells were stained LIVE for one hour with SPY555-actin in the medium, then fixed (obviously with PFA). Before fixing the SPY555 labelled of course the actin on the complete coverslip. The image shows a quarter of a coverslip (broken into pieces AFTER fixation, hence the SPY-staining only at the round edge.