I am attempting to perform some IHCs on Sprague-Dawley rat primary lymphatic endothelial cells, but I can't seem to get consistent results. When I attempt to image my cells after staining, the cells are either completely missing or are very scarce on the slide. If there happens to be any cells there, the signal is usually very weak. I'm using VE-cadherin, Claudin-5, Occludin, and ZO-1 antibodies to look at tight/adherens junction protein regulation.

Is anyone familiar with doing IHC on these types of cells? I'm thinking that my fixation method isn't working, but I'm pretty new to this technique so I might just be missing something!

My protocol is below for reference. I've tried fixing with 4% PFA and methanol separately and have gotten the same results.

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Perm buffer: 0.3% Triton X-100 in PBS

Blocking buffer: 1% Albumin, 2% normal donkey serum, and 0.3% Triton X-100, in PBS

1. Fix with 4% PFA for 30 minutes (I've also tried methanol at -20 C)

2. Wash with PBS. 2 x 5 min

3. Permeabilize tissue with 0.3% Triton X-100 in PBS for 30 min

4. Wash in PBS, 2 × 5 min

5. Block slides with blocking buffer for 1 hr at room temp

6. Prepare primary antibody in blocking buffer (1:200)

7. Incubate slides with primary at 4°C overnight

8. Wash slides in PBS

9. Prepare secondary antibody in blocking buffer 1:200 without triton

10. Incubate slides with secondary in the dark for 90 min at room temp.

- Minimize light after this point

11. Wash slides 2 × 5 min in PBS

12. Add DAPI (Prolong Gold) and coverslip

13. Allow to dry before imaging

Thanks for the help!!