I am trying double IF staining of parkin/COX IV or p62/COX IV for an autophagy study, I found either the signal was very weak (compared with the 2nd Ab only control, e.g. parkin or COX IV), or some non-specific dot signaling (eg. in the p62 staining). The cardiomyocytes were fixed with 4% paraformaldehyde at RT for 10-15 min, and permeablilized with 0.3% Triton X 100 (together with 5% NGS for blocking) at RT for an hour. I have tried increasing the 1°Ab concentration and incubating the samples with 1° Ab at RT for 1h or at 4°C overnight. There was no significant difference. I have also tried permeabilizing the cells with cold methanol for 10 min at -20C after paraformaldehyde fixation, there was not much improvement. By the way, what the pattern of parkin and p62 staining look like in normal primary cardiomyocytes. I would appreciate very much any input and help.
I have been having luck with dual label IF in adult rat cardiomyocytes, however not using the same primary abs as you. I am using estrogen receptors and Caveolin 3 primaries. The secondary abs I use are Cy3-conjugated donkey anti-rabbit IgG (H+L) (Jackson ImmunoResearch; 711-165-152) and Fluorescein (FITC)-conjugated donkey anti-mouse IgG (H+L) (Jackson ImmunoResearch; 715-095-150).
I use the following protocol-
Immunofluoresence Protocol
1. Cut 20 µm sections of frozen heart tissue (unfixed), mount on Polysine glass slides (ThermoScientific).
2. Post-fix tissue by placing drops of freshly made 4% PFA on sections. Incubate for 15 min at room temp. and wash 4X with 1XTBS for 5 min each. Circle around each section with wax pen.
3. Block the sections for 2 hr at room temp. (1X TBS, 0.2% TWEEN, 10% donkey serum, pH 7.4).
4. Prepare primary antibodies in 1X TBS, 0.2% TWEEN, pH 7.4 (TBST).
5. Incubate with primary antibody(s) (1:50-or what is recommended on product information sheet) for 24 hr in the cold room (in a humidity chamber). Make up enough ab dilution for ~100μl/section.
6. Wash the sections with 1XTBS for 10 min.
7. Wash 2X with TBST for 10 min each.
8. Prepare the secondary antibody(s) (1:150) in TBST in the dark.
9. Incubate the sections in the secondary antibody for 2 hr at room temp. in the dark.
10. Wash sections with 2X TBS + Triton (1X TBS + 0.1% Triton) for 5 min.
11. Mount the slides with Vectashield (Vectorlabs).
12. Leave the slides in a covered slide holder overnight before imaging with a confocal or fluorescence microscope. Fluorescence images should be obtained within 7 days after experiment.
Let me know if you would like any additional info.
-Rebecca
This was very similar to my Imunofluorescent Cytochemistry Protocol-
Immunofluorescent Cytochemistry
1. Aspirate media from the well and washed 3X in chilled PBS
2. Fix in 4% paraformaldehyde (30 min in cold room)
3. Wash in PBS (1x 5 min at r.t.)
4. Permeabilize cells with 0.2% Triton X-100 in PBS (10 min at r.t.)
5. Wash in PBS (3x at r.t.)
6. Block in 1 ml (1X TBS, 0.2% TWEEN, 1% BSA, 5% donkey serum, pH 7.4) in cold room.
Next day:
1. Incubate in primary antibody (1:50 in 1% BSA/TBS) – 2.5hrs in cold room.
2. Wash in TBS, 0.2% TWEEN (5 min 3X at r.t.)
3. Incubate in 2° (1:150 in blocking solution devoid of serum) – 1hr at r.t. in dark.
4. Wash in TBS (2x 5 min at r.t.)
5. Mount the cover slips on slides using Vecta shield (with DAPI)
6. Seal (always keep in dark)
You're welcome, I hope it helps. Yes, you can aliquot and store the 4% PF at -20.
Rebecca
For us permeabilising with 0.1% saponin in blocking buffer (PBS + 3% goat serum + 1% BSA) works well in primary cardiomyocytes. The protocol we usually follow is the following:
1.Fix cells in 4% PFA in PBS - 20 min at room temperature.
2. Wash once with 5 min incubation in PBS + 100 mM glycine to quench unreacted PFA.
3. Wash 3x PBS.
4. Block and permeabilise the cells for 45 min in 0.1% saponin in blocking buffer (PBS + 3% goat serum + 1% BSA).
5.Incubate overnight with primary antibody made in 0.1% saponin in blocking buffer.
6. Wash 5x with Wash buffer (0.1% saponin, 1% BSA in PBS).
7. Incubate 1h30 at RT with secondary antibody made in 0.1% saponin in blocking buffer.
8. Wash 5x with wash buffer.
9. Mount cells on coverslips with Vecta shield.
Please note that for visualising better some mitochondria associated proteins we had to combine 4% PFA fixing with -20oC methanol fix and permeabilisation and then again use 0.1% saponin in the incubation and washing steps.
I hope it helps.
Francoise
Thank you so much, Francoise. To clarify that for mitochondrial associated proteins which I am most interested in, how long do you incubate the cells with methanol at -20C? I assume that you fix the cells with 4% PF at RT for 20 min, then quench and wash, and then incubate the cells with methanol at -20C, right?
Li
So looking back at my lab book here are the steps we performed:
We did 4% PFA for 20 min at RT as usual and then we still quenched with 100 mM glycine in PBS. Next 5 min incubation with methanol at -20oC. After washing with PBS the blocking was performed only with blocking buffer as in my previous post but without saponin (sorry my mistake). The primary and secondary antibodies were incubated in blocking buffer without saponin. Washes with PBS-1%BSA.
Kind regards
Francoise
Thank you, Francoise. I have tired the similar method as I mentioned with my original question. However, I found that the signal of mitochondrial marker COX IV was only good at the location where it is close to the cytoplasmic membrane, e.g. at the confocal image layers where were generally close to the top, bottom or side of the cells. I am wondering whether it was due to the insufficient permeabilization? Did anybody have the similar experience? Any answers for that? Thanks!
Li
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