This problem was discussed recently. The main opinion was precipitate of plasmids with transfecting agent.

Just a few question to understand better:

- how many hours (or days) have passed after transfection prior you perfomed IF protocol?

- have you checked your DAPI batch?

- have you tried with HOECHST instead of DAPI? I do prefer H. instead of D. when I perform IF on HEK, because H. its a little bit cleaner.

- how do you use poly-lysine (how much and how long)?

It could also be a mycoplasma infection, which shows with DAPi staining as little specs around your cells. 

Have you analysed your samples for mycoplasma infection? If not, you should probably do it as it could also impact your research and make your results not reliable. There's PCR kits commercially available for this analysis.

Good luck

Daniil and Andrea's recommendation sound very reasonable.

1. You used too much DNA plasmid so that it disturb your IF result. When I do transfection, I only usse 500-600ng DNA plasmid for 4 of 96 wells. a)WHy dont you try to reduce your DNA concentration? b) Change the buffer for transfection. Using the buffer with lower salt concentration then. I often use the buffer from the transfection kit we bought from the company. I have no problem about this.

2. Hoetch is more sensitive than DAPI. You can try it.

3. Moreover, washing step should be increased and should be washed thoroughly. Increasing salt concentration is higly recommended to remove background of DAPI.

Trang

We've had this problem with other DNA stains and live cell imaging.  It is fairly easily fixed by a simple wash step prior to fixation.  I'll have to look up the buffer we used and get back to you.

Thank you so much Michael! Your help would be much appreciated.

Thank you all for the attentions. To answer Andrea and Trang, normally for a single well of a 8-well chamber slide I transfect no more than 200ng of plasmid DNA. At 24 h.p.transfection I wash the cells 3 times with PBS before fixation with 4%PFA. Normally I don't replace media after adding the transfection mixture so the lipid-DNA complex stays on cells for 24 hours (I did try replacing media at 6 h.p.t but no better results were obtained).

It sounds good. 

I agree with Michael, you will probably solve the problem by doing a more accurate washing step.

If I were in you, I would check my IF buffers for bacterial contamination: last month I noticed rod like spots near nuclei (hoechst staining); it turned out that one of my washing buffer was contaminated. I prepared it new and I did not have any spots.

Hi GuanQu.  I finally have that answer for you.  Sorry it took so long.  I haven't used this protocol, but found it for a lab mate who said it worked very well.  This is from Mirus Bio Technical Services if you ever want to call them fore more specifics.  Hope it helps.

Heparin Wash

The heparin wash procedure we use to remove/reduce transfection complexes is as follows. We use heparin sodium salt from porcine intestinal mucosa (Sigma cat # H3393). We simply weigh out the required amount and add 1x PBS to bring it to a 10 mg/mL stock and filter sterilize using a 0.2 mm filter (Steriflip, Millipore). Then we remove the medium from the cells and wash 3x with the solution, rocking the plate side-to-side and back-and-forth between washes. Then replace with normal growth medium.

Thank you so much Michael for the protocol. Just one more question what is the working concentration of heparin-PBS?

According to the protocol, 10 mg/ml in PBS.  Hope it works for you!

hello, I'm experiencing the same problem. How did you solve it? Thanks.