I have grown biofilm and I need to fix it and stain it.
1. Clean the entire disk carefully and aseptically with pipette so that extra biofilm which are present over the walls.
2. Cells are fixed for 20 minutes at room temperature with 4% paraformaldehyde in PBS followed by 2 washes with PBS to remove excess formaldehyde and stop the fixing reaction.
3. Biofilm on the cover glass are stained for 15 min with 3 microlitre mixture of both stains (Syto 13 and HI)
4. Cultures are kept in dark for 1 hr at room temperature.
Any suggestions for my protocol. I have attached the biofilm picture grown in the glass disk.
There are a few things I would suggest considering:
- fixation time of much longer than 20 minutes for biofilms. This will give the fixative time to actually cross link and fix the biofilm matrix. I would recommend at least overnight.
- Avoid using PBS to wash, or wash with PBS but then add water in place of PBS at the final step. PBS can interfere with fluorescence microscopy and the binding of fluorescently labelled stains.
Have you tried the stain/microscopy already?
Thank you Daniel for your suggestions. No I didnt try the stain/microscopy yet. I prepared the protocol so was looking for any suggestions. It would be really helpful to me.
Thanks
May I know what staining technique are you going to do with this biofilm? From my experience of doing FISH followed by confocal on mixed biofilms grown on cover slip in flow cell biofilms, you may need to pre-fix with 50% EtOH to ensure that your probes will pass through the cell wall and attached firmly on the target site. Attached is my protocol that my group have optimised for more than a year for your reference:
"At the completion of the 24 h incubation period, the biofilms were rinsed in situ with sterile PBS (Sigma-Aldrich, Australia) to remove non-adherent cells. Subsequently, the biofilms were fixed with 50% ethanol for 1 h at room temperature followed by washing with PBS for 30 min. The fixed biofilms were embedded in 20% acrylamide with 0.02% of ammonium persulfate and 0.8% of N,N,N,N tetramethylethylenediamine (TEMED), and incubated at room temperature for 30 min to solidify the gel. The coverslip was then removed and the biofilm embedded in the polymerised acrylamide slab was stored in PBS at 4 °C prior to fluorescent in situ hybridisation (FISH) staining."
In your case, you don't have to include ammonium persulphate and TEMED. Hope that helps a bit. Cheers.
What kind of CLSM do you used? We microscoped the biofilms directly after staining with an 96 well plate inlet (maybe you can use a 6 or 12 well plate if you have the inlet for the microscope?) so no fixation or anything was needed. Worked perfect. Because if you want to use LIVE/DEAD staining, i would dissuade you from using Ethanol or something like that because it will influence your results.
Daniel Morse Do you have a reference to support the claim "PBS can interfere with fluorescence microscopy and the binding of fluorescently labelled stains"? It would be very much appreciated.
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