Hi,

I am currently attempting to optimize an IHC protocol for some lymphatic endothelial cells that I'm studying. I've been having a lot of trouble getting consistently good images after staining, and am looking for any suggestions as to how I can fix this problem. I am staining for Occludin, and VE-cadherin tight junction proteins (images attached; VEC - red, Occ - Green).

Most of the images I get show very non-specific signal (I've attached some to this post so you can see for yourself). The proteins are extracellular membrane proteins, but I haven't been able to see that yet. My protocol is here for reference:

*All washing steps were done by gently pipetting solution onto walls of each well instead of directly onto cells - this prevents cell detachment

*Antibody wash solution: 0.1% BSA (in PBS)

*Blocking Solution: 10% Normal Donkey Serum

1. After treatment of cells on microscope slides, wash cells 3x, 5min with PBS.

2. Add 4% PFA (150 uL/well if 8-well slides are used). Let sit for 15 min.

3. Remove PFA and wash 3x, 5min each with PBS

4. After last PBS wash, remove PBS and block with antibody blocking solution for 30 min

5. Remove blocking solution and then add primary antibody diluted in blocking buffer for 1 hr in 4 degrees C

• 1:200 concentration for all ABs

6. Remove primary AB and wash 3x, 5min w/ AB wash solution

7. Add secondary AB diluted in blocking buffer for 1 hr; dark room, room temp

- I typically do FITC and TRITC conjugated secondaries

8. Remove secondary and wash 3x, 5 min with antibody wash

9. Add prolong gold with DAPI to each slide and add cover slip; seal edges

10. Image slides

Thanks in advance for reading and for the help!!